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Search Fundamentals of Biochemistry RNA: Structure and Function Ribonucleic acids are very similar in chemical structure to DNA except they contain ribose instead of deoxyribose. They also have the pyrimidine base uracil instead of thymine, as shown in Figures 1 and 2 above. These two small changes (but mostly the first) confer on it a very different set of biological functions than DNA. This should not surprise us and the basis of all chemistry and biochemistry is that chemical structure determines chemical and biochemical functions and activities. We discussed in the previous section how RNA can adopt complex tertiary structures, which requires the presence of more noncanonical base pairs and chemical modification of bases. In this section we wi,ll explore the plethora of different types of RNA structures and their functions. The sequence of RNA is made from DNA through a process called transcription (converting the information of DNA, a nucleic acid, into RNA, another nucleic acid). RNA can form double-stranded helices but typically these are viral in origin. DsRNA is a pathogen-associated molecular pattern (PAMP) that binds Toll-like receptor 3 (TLR3) as we saw in Chapter 5.5. If both strands of DNA are transcribed, the resulting strands can anneal form dsRNA. In addition, a single strand of RNA can fold on itself if the 5' and 3' ends are complementary to form a stem-hairpin loop. Figure \(1\) shows a stem-loop from a messenger RNA (4QOZ) when it is bound to a specific RNA binding protein (not shown). Larger ss-RNA can form tertiary structures with many regions of intrastrand hydrogen bonds forming secondary structures, as shown in Figure \(2\) for one type of RNA called a transfer RNA Figure \(3\) a computed model for secondary structure within a much larger single RNA molecule S11 that is part of the ribosome. The figure shows color-coded differences in accessibility when the S11 RNA is free (blue) and bound to the protein NSP2 (red), which induces structural rearrangements. You can imagine that a different set of intrachain H-bonded double-stranded regions could easily form, with the most likely determined by sequence, local environment, and protein binding partners. Each RNA molecule would have a thermodynamic folding landscape similar to protein. Programs are available to determine secondary structures from RNA sequences. Each RNA molecule would have a thermodynamic folding landscape similar to protein. Also, the structures are dynamic as we saw with proteins. Another feature that makes RNAs complicated is that many different types of RNA are made from DNA using RNA polymerases. They are loosely divided into two types of RNA. One is coding RNA, which contains the sequence information that will be translated into a protein sequence. The other type is called noncoding RNA. These RNAs regulate a myriad of cellular processes including transcription to produce the coding RNA. Coding RNA The DNA template from which the coding sequence of a translatable RNA is produced is called a gene. The coding RNA which has the exact sequence that is translated into protein is called messenger RNA (mRNA). The exact sequence of RNA in mRNA that encodes a protein is derived from a longer contiguous DNA sequence in the nucleus from which sections called intervening sequences or introns have been removed. The coding sequences of DNA which are separated by introns are called exons. When coding DNA is transcribed, a long contiguous sequence containing both exons and introns is transcribed into one long primary transcript called heteronuclear RNA. The introns in the heteronuclear RNA are removed, in a splicing reaction catalyzed by a large complex called the spliceosome to form mRNA. The process is illustrated in Figure \(4\). The first RNA sequence made is the heteronuclear RNA. The long single stranded mRNA molecule binds to ribosomes, nanomachines which orchestrate the translation of the mRNA sequence into a protein sequence. There are around 20,000 human genes that produce an even large number of mRNA that arise from differential splicing of the primary transcript. Noncoding RNA (ncRNA) Not long ago, few thought about possible RNA transcripts from non protein-coding regions of the genome, except for two types of RNA required for the translation of mRNA. These two are ribosomal RNAs (rRNA) which are found in ribosomes and transfer RNAs, to which amino acids are esterified and transferred to a growing protein chain of the ribosome. Many more classes have been discovered and given names that are quite confusing to someone more vind with protein structures. One way to classify noncoding RNAs (ncRNAs), which implies non-protein coding RNAs is based on size. • short noncoding RNAs (sncRNAs) are <200 nucleotides • long noncoding RNAs (lncRNAs)are >200 nucleotides These function to regulate gene expression at both the transcription and post-transcriptional levels. Some have catalytic functions. Some affect chromosome structure and chemical modification. Long Noncoding RNAs (lncRNAs) There many be between 16,000 to over 100,000 human lncRNAs encoded into the genome which adds much complexity to our understanding of the function of RNA transcripts. An online lncipedia is a database of searchable lncRNA sequences. There are many types of lncRNAs. The first we will consider is ribosomal RNA. a. Ribosomal RNA (rRNA): These RNAs fit the simple definition of lnRNAs (>200 nucleotides and are not protein- ing), but most would not think of them as lncRNAs since they have always been in their own category of a nonprotein-coding gene. rRNAs vary in length from between 1500 and 3000 nucleotides long in bacteria and about 1800 and 5000 nucleotides long in humans and are the core structure of ribosomes, the nanomachines which translate bound mRNA into a protein sequence. Figure \(5\) shows an interactive iCn3D model of the structure of 23S rRNA of the large ribosomal subunit from Deinococcus radiodurans (2O44) (long load time). The red (highlighted yellow) spacefill is the 5' start of the rRNA. The chain has a complex tertiary structure, much like a protein sequence, and ends at the cyan spacefilling 3' end. It has 2880 nucleotides. Let's focus on more classical examples of long noncoding RNAs (i.e not rRNA). One way to categorize them is based on the position in the genome that encodes them. The different types include long intergenic noncoding RNAs (lincRNA), intronic lncRNAs, antisense RNAs (as lncRNAs) and other variants. These are illustrated in Figure \(6\), where the lncRNA is shown in pink. Another variant is exon and intron-containing circRNAs (EIciRNAs) as illustrated in panel B in Figure \(13\). These are presumably produced from pre-mRNA for a given mRNA, and appear to regulate gene expression through RNA-RNA interactions with U1 snRNA, which starts the assembly of the spliceosome on pre-mRNA when it binds to the 5′ pre-mRNA splice site. Now let's consider more typical examples of long non-coding RNAs (lncRNAs), which are often bound to target proteins. b. mamRNA (a lnc RNA) The lncRNA named mamRNA (Mmi1 and Mei2-associated RNA) binds two proteins, Mmi1 and Mei2 in Schizosaccharomyces pombe that control the balane between meisois and mitosis in yeast. (Schizosaccharomyces pombe is a "fission" yeast that divides by fission and not budding.) The MamRNA has two variants of length 550 and 700 nucleotides. Binding of mamRNA leads to the ubiquitinylation of the Mei2 in the complex. Mmi1 R is an RNA-binding protein that binds to a modified version of adenosine that has been methylated at N6 and is found internally in mRNA. Mei2 (meiosis protein 2) is necessary of meiosis. The binding of mamRNA leads to the ubiquitinylation of the Mei2 in the complex, targeting it for proteolysis. Mei2 concentrations relatively increase, shifting yeast from mitosis to meiosis. Figure \(7\) shows a cartoon depicting these interactions. Figure \(8\) shows an interactive iCn3D model of the S. pombe Mei2 RRM3 protein domain bound to the Mei2 binding "domain" of mamRNA (6YYM) which in this structure is only 8 nucleotides long (not the full length this lncRNA which is 550 and 700 nucleotides long). c. ToxI - a lnRNA inhibitor of the endonuclease ToxN Those with a more chemistry-centric background might be surprised to know that viruses also "infect" bacteria. These viruses are called bacteriophages. It is estimated that there are over 1030 in nature. Some covalently incorporate into genomes where they reside and are incorporated permanently into the genome. They are a main driver of bacterial genome evolution as they shape the bacteria's immune response and adaptation. One very interesting example is the type III toxin-antitoxin (TA) system in E. Coli. It consists of a toxin, ToxN, which is a nuclease that cleaves internally after the second A in a AAA sequence. It acts on mRNA but especially pre-mRNA sequences. It is inhibited by the binding of a lncRNA called ToxI (toxin inhibitor). The RNA sequence of the ToxI inhibitor has 36 "domain" repeats of a pseudoknot, one of which is sufficient to inhibit the ToxN. The ToxN endonuclease cleaves the ToxI lncRNA as it assembles the complex. It also cleaves its mRNA. Figure \(9\) shows an interactive iCn3D model of the protein toxin (ToxN):lncRNA (ToxI) which is a shortened version of 29 nucleotide section from Pectobacterium atrosepticum (2xdb). Short Noncoding RNA Short noncoding RNAs (sncRNAs) are less than 200 nucleotides in length. By definition, this would include transfer RNAs (tRNAs) which bring to the ribosome amino acids covalently attached to their 3' end for incorporation into a growing protein chain during the translation of mRNA. As with rRNA for lncRNAs, these are really in a class of their own. Others include small nuclear RNAs (snRNAs) involved in splicing, small nucleolar RNAs (snoRNAs) involved in the modification of rRNAs, and microRNAs (miRNAs), involved in the inhibition of translation and transcription, PIWI-interacting RNAs (piRNAs), and endogenous small interfering RNAs (siRNAs). It is difficult to remember the subtle difference among these, which makes them a bit difficult to understand. We will tell their stories with a few targeted examples. a. Transfer RNA: Transfer RNAs act as adapter molecules between transcription and translation. They are between 76 and 90 nucleotides long and have a cloverleaf shape. An enzyme, aminoacyl-tRNA synthase, covalently attaches a select amino acid at its 3' end. Another end of the tRNA hydrogen bonds through 3 nucleotides (the anticodon) to a triplet nucleotide (the codon) on the mRNA that encodes a specific amino acid at that triplet position. Figure \(10\) shows an interactive iCn3D model of the structure of yeast phenylalanine tRNA (1EHZ) b. Small nuclear RNA (snRNA): The spliceosome is a nanoparticle that catalyzes the removal of introns from pre-mRNA in eukaryotes (prokaryotes appear devoid of introns). The yeast spliceosome has a molecular weight of 1.3 million and contains 5 small ribonucleoproteins (RNPs) with many other associated proteins. Each of the 5 RNPs has a small nuclear RNA (U1, U2, U4, U5 and U,6) which is enriched in uracils. U6 is highly conserved and is directly involved in catalysis. Figure \(11\) shows an interactive iCn3D model of the core structure of the U6 small nuclear ribonucleoprotein complex with most of the U6 RNA bound. c. MicroRNAs (miRNAs) and small inhibitory RNAs (siRNAs) MicroRNAs (miRNAs) control the expression of thousands of genes in plants and animals. They are single-stranded but fold on themselves to form a stem-hairpin. The miRBase is a microRNA database containing almost 40,000 miRNA sequences. miRNAs are highly conserved and are found in animals, plants, and some unicellular eukaryotes. They interact with the 3′ untranslated regions of mRNAs and inhibit or prevent their translation. Several key proteins, RNA polymerase II, Drosha and Dicer are involved in the canonical pathway while the others appear to be independent of Drosha, which is a ribonuclease III double-stranded (ds) RNA endoribonuclease. Dicer is a dsRNA) endoribonuclease which cleaves long dsRNAs and short hairpin pre-microRNAs (miRNA) into fragments of either 21-23 nucleotides (short interfering RNA) or 19-25 nucleotides (microRNAs). Each has two nucleotides that are unpaired at the 3' end. These bind to the enzyme complex RISC ( RNA-induced silencing complex) whicn then targets the to mRNA complementary to the siRNA/miRNA (RISC) causing cleavage of the mRNA and hence inhibiting translation. Small inhibitory RNAs (siRNAs) are very similar to miRNA (to the point that differentiating between them is somewhat arbitrary). They both engage in RNA interference (RNAi) of mRNA translation. Here some reported differences: • The substrate for dicer cleavage is dsRNA (that could be added exogenously) of length 30-100+ for siRNA but the actual pre-miRNA of length 7-100 nucleotides that may contain hairpins with some mismatches (i.e. not a perfect stem and hairpin) • The final RNA after dicer processing is double-stranded for both and 21-23 nucleotides long for siRNA and 19-25 for miRNA • siRNAs are perfectly complementary to the target mRNAs while miRNAs, which are not necessarily perfectly complementary, bind typically to the 3' untranslated end of the mRNA • Because of the perfect complementarity to target mRNA, siRNA interact with only one mRNA while miRNAs, given that they are not perfectly complementary to their target sequences, can bind different mRNAs • Given their higher affinity binding, the siRNA leads to dicer endonuclease cleavage of the target mRNA while inhibition of mRNA translation by miRNAs arises from binding of the miRNA to the mRNA or, if the match between the miRNA and mRNA is high enough, endonuclease cleavage of the mRNA. Figure \(12\) shows a canonical and several alternative pathways for their transcription and processing from the noncoding miRNA genes. Figure 1. Canonical and non-canonical pathways of microRNA biogenesis. (A) Canonical pathway—microRNA gene is transcribed by RNA polymerase II into primary microRNA (pri-miRNA), cleaved by microprocessor complex Drosha/DGCR8, and precursor microRNA (pre-miRNA) is exported from the nucleus to the cytoplasm by Exportin 5 (XPO5) and further processed by Dicer and its partners into 18–25 nucleotide long microRNA duplex with 2-nucleotide 30 overhangs. The guide strand is subsequently bound by the Argonaute proteins 1-4 (AGO1-4) and retained in the microRNA-induced silencing complex to target mRNAs for post-transcriptional silencing. (B) Mirtrons—generated through mRNA splicing independently of Drosha-mediated processing step. (C) Small nucleolar RNA-derived microRNAs—Drosha-independent pathway. (D) Exportin 5-independent transport of pre-miRNAs from the nucleus to the cytoplasm has been described in the case of miR-320 family. (E) Dicer-independent processing of miR-451—pre-miR-451 is directly loaded into AGO2, cleaved and trimmed by poly(A)-specific ribonuclease PARN to produce mature miR-451. Gregorova et al. Cancers 2021, 13, 1333. https://doi.org/10.3390/cancers13061333. Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/) siRNA that are perfect matches to specific mRNA can be easily designed and purchased for translation inhibition and through that gene silencing studies. Both miRNAs and siRNA are potentially therapeutic as it is much simpler to design a drug that targets a mRNA sequence (a 1D sequence target) than a protein active site (a 3D target). Additionally, they can be used to inhibit protein synthesis of target proteins that don't have a "druggable" active site. The protein Argonaute is involved in miRNA and siRNA silencing of genes through their mRNAs in the RISC ( RNA-induced silencing complex). RISC contains the protein argonaute 2 (AGO2) bound to a "guide" RNA which is either microRNA (miRNA) or short interfering RNA (siRNA). It is the miRNA or siRNA that directly interacts with the "target" - the mRNA. Example miRNA: Figure \(13\) shows an interactive iCn3D model of human Argonaute2 Bound to a Guide (miRNA) and Target RNA (4W5O). The two RNA sequences are 5' UUCACAUUGCCCAAGUCUUU 3' and 5' CAAUGUGAAA 3'. Example: siRNA (Small interfering RNA) Virus genomes are ultimately decoded into new viruses by the host replication, transcription and translation machinery. Host cells have evolved ways to silence mRNAs from viruses. Unfortunately, viruses, in response, evolve ways to suppress host RNA silencing. Many viral proteins are used to suppress silencing by the host. One is the viral p19 protein, which preferentially binds to host short interfering RNAs (siRNAs) than to microRNAs (miRNAs). A single mutation in the viral p19 proteins changes selectivity which allows it to bind to a specific human miRNA called miR-122. This shows the subtle complexities of protein:RNA interactions. Figure \(14\) shows an interactive iCn3D model of the viral suppressor of RNA silencing protein and a 21 residue small interfering RNAs (6BJV) Example: piRNA (a specific miRNA) Piwi proteins are RNA-binding proteins in plants and animals and are similar in structure to argonaute. They bind a guide RNA called piwi-interacting RNAs (piRNAs) and lead to the silencing of sequences in the genome called transposable elements that can move around the genome. piWi has endonuclease activity and can cleave mRNA. The piRNAs are just one type of miRNA. Figure \(15\) shows an interactive iCn3D model of Ephydatia fluviatilis (a sponge) PiwiA with a guide (piRNA) and-target RNA(7KX9) lncRNAs and miRNAs in the brain After the decoding of the human genome, many have been struggling to understand how the complexity of the human brain (large size, greater connectivity among neurons) arises given that we appear to have only around 20,000 proteins genes encoded by the genome (not counting small proteins of less than 100 amino acids). Long noncoding RNAs (lncRNAs) and miRNAs appear to be significant pieces of this puzzle. Their ability to regulate transcription during development may hold the key. Additional roles on these RNAs outside of transcriptional regulation are being discovered. Some are transported away from the nucleus to serve other functions in axons, dendrites, etc. For example, the lncRNA Gm38257 binds to proteins that structure the synapse. something rather unexpected: Instead of simply regulating gene expression, it binds to proteins The repertoire of miRNA appears to be significantly increased in "intelligent" organisms such as humans and octopi. For example, a large increase (179) in miRNAs occurs in proceeding in the evolutionary scale from mice (which have about 24,000 protein-encoding genes) to humans (around 20,000). miRNAs and lncRNAs may be involved (causative or correlative?) with brain disease. An example is the miR-124 is significantly elevated (3.5X) times higher in hippocampal cells from mouse models of Alzheimer's compared to normal mice. Altered expression of the lncRNA named Gomafu, RNCR2 or MIAT) appears to affect certain psychiatric diseases. f. small nucleolar RNA The nucleolus is a small structure in the nucleus that helps assemble the ribosomal RNAs that are synthesized in the nucleus. They then are transported through the nuclear membrane into the cytoplasm where they combine with proteins translated from mRNA in the cytoplasm to form complete ribosomes. As we will describe below, rRNA is chemically modified by enzymes (much like the post-translational modification of proteins). One such modification is 2'-O-methylation in archaea and eukaryotes. A class of small nucleolar RNAs (snoRNAs) which vary from 10-21 base pairs are called C/D RNAs, and they "guide" the modification. Hence they are also called "guide" RNAs. These snoRNAs bind to 3-4 proteins into ribonucleoproteins. Figure \(16\) shows an interactive iCn3D model of the box C/D ribonucleoprotein 40 nt snoRNA "guide" and a 10 nucleotide RNA target substrates. It appears that the maximal duplex RNA formed (from the guide and target) is 10 base pairs long.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/08%3A_Nucleotides_and_Nucleic_Acids/8.02%3A_Nucleic_Acids_-_RNA_Structure_and_Function.txt

Search Fundamentals of Biochemistry Now that we have an understanding of the structures of DNA and the structures and various functions of RNA, we can now more fully explore how their chemical similarities and difference contribute to different functions. Chemical modifications of DNA and RNA Post-translation modifications of proteins alter their structural/functional properties. Likewise, intentional chemical modifications of nucleic acid bases alter both their structures and potentially their transcriptional and translational status. Figure \(1\) shows common modifications of bases in DNA. Likewise, RNA is chemically modified. Figure \(2\) shows common modifications of bases in RNA. Methylation and subsequent hydroxylation to hydroxymethyl are common to both DNA and RNA. Methylation of DNA often represses the transcription of the DNA into RNA. Hence it has huge potential to alter gene transcription. Such changes to the DNA are called epigenetic modifications. These changes can be passed down to future generations as well and affect the phenotype of a cell. Histone proteins involved in DNA packing into nucleosomes can also be methylated and acetylated, altering the interaction of the DNA with the nucleosome core and further packing, again affecting transcription. The chemical modification to RNA also can change the reading out of the genome. The epitranscriptome refers to the collective chemical modifications to RNA, and its understanding is part of new field called epitranscriptomics. Mutations Mutation can arise from the chemical modification of bases. Uracil in RNA is a demethylated form of thymine in DNA. In RNA, AU base pairs replace AT base pairs. Why the need for uracil in RNA? The question could be rephrased as to why the need for thymine, with its extra methyl group, in DNA. It's useful to think about the consequence of replacing a single H in a molecule with a -CH3. Take HOH, water, as an example. Our bodies are over 60% water. We drink liters of water of concentration 55 M each day. Yet if we drink 0.07 L of methanol, CH3OH, half of us would die! Let's probe some consequences of the U (no -CH3) and T (with -CH3) changes in DNA. It can get confusing but just remember that the normal base pairs in DNA are AT, but AU base pairs also form (they norm in RNA). The -CH3 substituent on thymine does not affect its base pairing. a. Spontaneous deamination of cytosine in DNA Why are we now discussing cytosine in DNA? One reason is that the most common mutation in DNA is a C to T replacement. One way that happens is through the spontaneous hydrolytic deamination of cytosine in DNA to uracil, which we have presumed to be found only in RNA. The mechanism for this deamination and subsequence conversion of a GC to an AT base pair is shown in Figure \(3\). The inset box shows a simplified mechanism for spontaneous deamination. Hence a possible consequence of the deamination reaction is a GC to AT base pair mutation if the uracil in DNA is not removed before DNA replication. Fortunately, the enzyme uracil-DNA glycosylases can remove any uracils found in DNA, leaving an abasic site, which can be fixed with DNA repair enzymes. We can now ask the question, why T and not U in DNA? Pretend you are a DNA repair enzyme and you see a UA base pair in DNA. How can you tell if the UA base pair is correct and intended to be there or if it should be a CG base pair that underwent deamination? The most common uracil-DNA glycosylases remove the uracil whether it is across from guanine, the correct base but which can not hydrogen bond with uracil (in the green oval in Figure \(3\)), or if is across from adenine, the wrong base (in red oval), which is present after a round of replication. Evolution has addressed this problem by adding a methyl group to uracil to form thymine and using that base, which forms a base pair with adenine. Now no decision on which base across from a uracil (guanine if the uracil arose from deamination) or across from a "uracil-like" thymine (adenine) is correct. b. Other mutations Since we are considering chemical modifications to DNA and mutations, it is appropriate to give a more expanded background on them. In addition to mutations caused by spontaneous hydrolytic deamination of cytosine, mutations can also arise through the addition of a wrong base during DNA replication, by chemical damages caused by radiation or chemical modifying agents. How many mistakes in replication are made? If you received a 99% on an examination, you would be ecstatic. That's not good enough for DNA replication. In Cell Biology by the Numbers, they calculate it this way. Assume the replication /repair is so good that it takes 108 replications to make a mistake (an error rate of 10-8/BP). Assume also there are 3 x 109 base pairs in the human genome. This leads to a mutation rate 10-100 mutations/genome/generation or about 0.1-1 mutations/genome/replication. Not bad! Figure \(4\) shows how common point mutations might arise just randomly. Chemical agents also can cause point mutations. Figure \(5\) shows point mutations arising from oxidative deaminations (not hydrolytic) by nitrous acid/nitrosamines and from alkylating agents. Figure \(6\) shows a variety of alkylating agents with mutagenic potential. Finally, large-scale changes in chromosome structure can also occur as shown in Figure \(7\), usually with profound consequences. Why DNA and RNA - A chemical perspective Asking a "why" question (like above) in the sciences is really not appropriate as such teleological questions are more philosophical or religious. Yet we will in this section, in part, to be in the company of Alexander Rich, who wrote a very cool article entitled "Why RNA and DNA have different Structures". Given that RNA expresses catalytic activities and can carry genetic information (some viruses have ds and ss RNA as their genome), it has been suggested that early life might have been based on RNA. DNA would evolve later as a more secure carrier of genetic information. An inspection of the chemical properties of DNA, RNA, and proteins shows them to have attributes needed for their expressed function. Let's examine each for structural features that might be important for function. a. Why does DNA lack a 2' OH group (found in RNA), which has been replaced with hydrogen? This required the evolutionary creation of a new enzyme, ribonucleotide reductase, to catalyze the replacement of the OH in a ribonucleotide monomer to form the deoxyribonucleotide form. One possible explanation is offered in the figure below. DNA, the main carrier of genetic information, must be an extremely stable molecule. An OH present on C'2 could act as a nucleophile and attack the proximal P in the phosphodiester bond, leading to a nucleophilic substitution reaction and potential cleavage of the link. RNA, an intermediary molecule, whose concentration (at least as mRNA) should rise and fall based on the need for a potential transcript, should be more labile to such hydrolysis. Figure \(8\) shows a possible reaction diagram for the internal cleavage of RNA. (The reaction would probably proceed with no actual intermediate, but just a transition state. b. Why do both DNA and RNA contain a phosphodiester link between adjacent monomers instead of more "traditional" links such as carboxylic acid esters, amides, or anhydrides? One possible explanation is given below. Nucleophilic attack on the sp3 hybridized P in a phosphodiester is much more difficult than for a more open sp2 hybridized carboxylic acid derivative. In addition, the negative charge on the O in the phosphodiester link would decrease the likelihood of a nucleophilic attack. The negative charges on both strands in ds-DNA probably help keep the strands separated allowing the traditional base pairing and double-stranded helical structure observed. The cleavage of the phosphodiester link in DNA and a hypothetic ester link is shown in Figure \(9\). Again, the reaction of the phosphodiester shows a pentavalent intermediate, but most like the reaction proceeds directly from the transition state. c. Why is DNA found as a repetitive double-stranded helix but RNA is usually found as a single-stranded molecule that can form complicated tertiary structures with some ds-RNA motifs? Another reason for the absence of the 2' OH in DNA is that it allows the deoxyribose ring in DNA to pucker in just the right way to sterically allow extended ds-DNA helices (B type). The pucker in deoxyribose and ribose can be visualized by visualizing a single plane in the sugar ring defined by the ring atoms C1', O, and C4'. If a ring atom is pointing in the same direction as the C4'-C5' bond, the ring atom is defined as endo. If it is pointing in the opposite direction, it is defined as exo. In the most common form of double-stranded DNA, B-DNA, which is the iconic extended double helix you know so well, C2' is in the endo form. It can also adopt the C3' endo form, leading to the formation of another less common helix, a more open ds-A helix. In contrast, steric interference prevents ribose in RNA from adopting the 2'endo conformation, and allows only the 3'endo form, precluding the occurrences of extended ds-B-RNA helices but allowing more open, A-type helices. Figure \(10\) shows another comparison between the A-RNA and B-DNA double helices and the C'3 and C'2 endo forms of the ribose Figure \(5\) shows interactive iCn3D models of the pentoses in a strand of A-RNA (413D), double-stranded, left, and B-DNA (1BNA), double-stranded, right. C'3-endo ribose, A-RNA (413D, double stranded) C'2 endo ribose, B-DNA (1BNA, double stranded) Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...KPueqrBADczh26 Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...BEn5nqsCQG2JH6 d. What about the molecular dynamics of A-RNA and B-DNA? The information above suggests that the sugar ring of DNA is conformationally more flexible than the ribose ring of RNA. This can be inferred from the observation that dsDNA can adopt B and A forms, which requires a switch from the 2' endo in the B form to the 3'endo form in the A form. The smaller H on the 2'C would offer less steric interference with such flexibility. The rigidity in ribose is associated with a smaller 5'O to 3'O distance in RNA leading to a compression of the nucleotides into a helix with a smaller number of base pairs/turn. The increased flexibility in DNA allows rotation around the C1'-N glycosidic bond connecting the deoxyribose and base in DNA, allowing different orientations of AT and GC base pairs with each other. The normal "anti" orientation allows "Watson-Crick" (WC) base pairing between AT and GC base pairs while the altered rotation allows "Hoogsteen" (Hoog) base pairs. The different orientations for an AT base pair are shown in Figure \(11\). The Watson-Crick (WC) and Hoogsteen (HG) base pairs in B-DNA are in a dynamic equilibrium with the equilibrium greatly favoring the WC form as indicated by the arrows in the figure above. In a DNA:protein complex, the WC ↔ HG equilibrium can favor the WG form for AT and GC+ forms (in the latter, the C is protonated) when those base pairs are also involved in protein recognition. They can also occur more frequently in damaged DNA. In contrast, molecular dynamic studies show that the HG base pairs A-U and GC+ are strongly disfavored in ds A-RNA. One type of DNA damage is methylation on N1-adenosine and N1-guanosine. This modification prevents normal Watson-Crick base pairing but for DNA, these modified bases can still engage in Hoogsteen base pairing, preserving the overall structure of dsDNA and its ability to stably carry genetic information. This same methylation occurs normally in post-transcriptional modified RNA. Hence, N1 adenosine and N1 guanosine methylation prevent any type of base pairing in the modified RNA. These properties make DNA a better carrier of molecular information and offer another way to regulate the structural and functional properties of RNA. Hoogsteen base pairs can be found in distorted dsDNA structures (caused by protein:DNA interactions) but also in normal B-DNA. Figure \(12\) shows a Hoogsteen base pair between dA7 and dT37 in the MAT α 2 homeodomain:DNA complex (pdb 1K61). Note that the dA base in the Hoogsteen base pair is rotated syn (with respect to the deoxyribose ring) instead of the usual anti, allowing the Hoogsteen base pair. A Structural Comparison Now let's review the kinds of structures adopted by the 3 major macromolecules, DNA, RNA, and proteins. DNA predominately adopts the classic ds-BDNA structure, although this structure is wound around nucleosomes and "supercoiled" in cells since it must be packed into the nucleus. This extended helical form arises in part from the significant electrostatic repulsions of two strands of this polyanion (even in the presence of counter-ions). Given its high charge density, it is not surprising that it forms complexes with positive proteins and does not adopt complex tertiary structures. RNA, on the other hand, can not form long B-type double-stranded helices (due to steric constraints of the 2'OH and the resulting 3'endo ribose pucker). Rather it can adopt complex tertiary conformations (albeit with significant counter-ion binding to stabilize the structure) and in doing so can form regions of secondary structure (ds-A RNA) in the form of stem/hairpin forms. Proteins, with their combination of polar charged, polar uncharged, and nonpolar side chains have little electrostatic hindrance in the adoption of secondary and tertiary structures. RNA and proteins can both adopt tertiary structures with potential binding and catalytic sites, making them ideal catalysts for chemical reactions. RNA, given its 4 nucleotide alphabet, can also carry genetic information, making it an ideal candidate for the first evolved macromolecules enabling the development of life. Proteins with a great abundance of organic functionalities would eventually supplant RNA as a better choice for life's catalyst. DNA, with its greater stability, would supplant RNA as the choice for the main carrier of genetic information (Figure \(13\)): A final note on the simplicity of the dsDNA structure. A mutation causing a single base pair change in DNA does not change the iconic ds-stranded DNA structure. If it did, DNA would not be a reliable molecule to store and read out the genetic blueprint. In contrast, a single mutation in the DNA leading to a single amino acid substitution may lead to a protein with altered structure and function. On one hand that could be deleterious or even fatal to the organism. On the other hand, the new protein structure might have new functionalities that allow adaptation to new environments or allow new types of reactions. Evolution would favor the latter.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/08%3A_Nucleotides_and_Nucleic_Acids/8.03%3A_Nucleic_Acids_-_Comparison_of_DNA_and_RNA.txt

Search Fundamentals of Biochemistry Chromatin When stained and viewed in a microscope, eukaryotic nuclear DNA in nondividing cells is observed in two different states, heterochromatin (dark areas) and euchromatin (light areas), as shown in Figure \(1\). The heterochromatin is darkly stained and found along the inner side of the nuclear envelope as well as inside the nucleus. Euchromatin doesn't stain well since it is more dispersed, and is found throughout the nucleus. The image above is of a cell in interphase, a part of the cell cycle in which the cell is in-between cell divisions. Here are some differences between heterochromatin and euchromatin: Heterochromatin: • contains genes that are transcriptionally inactive and not expressed; • is not found in prokaryotic cells; • in humans, one of the X chromosomes is silenced in heterochromatin while the other is transcribable and in euchromatin; • protects the tightly packed in it from nucleases. Euchromatin: • contains about 90% of genome DNA; • contains chromosomes that are unfolded into nucleosomal DNA resembling beads on a string and is characterized by looser association with packaging histone proteins; • is the only form of DNA in bacteria; • is transcriptionally active so it is open to the binding of RNA polymerase and transcription factors; • has some genes which can be transcriptionally silenced by moving them into heterochromatin. Look as long as you wish but you won't see the iconic pictures of chromosomes in Figure \(1\). They are there but dispersed. They are only discretely visible at certain times in the cell cycle when cells are preparing to divide. Cell Cycle For those with a more chemistry-centric background, a short review of the cell cycle would be helpful. Somatic (non-germ cells) spend only part of their lives preparing for and dividing into two cells in a process called mitosis. Figure \(2\) shows a simple model for the cell cycle. Before a cell gets ready to divide, it is in interphase (between mitotic phases), which encompasses the G1, S, and G2 phases of the cell cycle. In interphase, cells grow and replicate their DNA Mitosis or cell division occurs after G2 and consists of a series of new discrete phases, including prophase, metaphase, anaphase, and telophase, after which the cell divides in a process called cytokinesis. Table \(1\) below shows the stages of mitosis and cytokinesis which occurs after interphase. Let's start with a discussion of the structure of chromosomes as classically observed in mitotic cells. Then we will look more closely at the seemingly amorphous and more complicated structures of chromatin. Chromosomes Within eukaryotic cells, DNA is organized into long linear structures called chromosomes. A chromosome is a "thread-like" structure in the nucleus of animal and plant that consists of a single but long molecule of double-stranded DNA (so it's two ss-DNAs) with a myriad of bound proteins. The proteins bind to and condense the DNA molecule to prevent it from becoming an unmanageable tangle. Before typical cell division (mitosis), these chromosomes are replicated by DNA polymerase to make two identical chromosomes, one for each future daughter cell. The two identical chromosomes, called sister chromatids, bind to each other at a common structure called the centromere. A replicated chromosome (sister chromatids) bound to each other at the centromere, is shown in Figure \(3\). Each chromosome in the sister chromatid structure represents one chromatid. In humans, each cell normally contains 23 pairs of chromosomes, for a total of 46. In twenty-two of these pairs, called autosomes, each member of the pair is similar. The 23rd pair, the sex chromosomes, differ between males and females. Females have two copies of the X chromosome, while males have one X and one Y chromosome. Figure \(4\) shows a DNA karyotype with 22 pairs of autosomes and one pair of sex chromosomes. Karyotypes are prepared using standardized staining procedures that reveal characteristic structural features for each chromosome, usually from white blood cells. The karyotype of human cells shown in Figure 4 below shows an extra copy of chromosome 21 (trisomy 21), which causes Down's Syndrome. Each species has a unique chromosomal complement. For example, chickens have 39 pairs of chromosomes (78 total) with 38 autosomal pairs and one pair of sex chromosomes (Z and W). In this species, ZW chickens are female and ZZ chickens are male. The total length of the human genome is over 3 billion base pairs. The total length of the human genome is over 3 billion base pairs. The genome also includes mitochondrial DNA. Eukaryotic organisms (animals, plants, fungi, and protists) store most of their DNA inside the cell nucleus as linear nuclear DNA. However, DNA in the mitochondria and chloroplast are circular (with no ends). Bacteria and Archaea cells do not have organelle structures and thus, store their DNA only in a region of the cytoplasm known as the nucleoid region. Prokaryotic chromosomes consist of double–stranded circular DNA. Packing of DNA: Supercoiling The genome of a cell is often significantly larger than the cell itself. For example, if the DNA from a human cell containing 46 chromosomes were stretched out in a line, it would extend more than 6 feet (2 meters)! How is it possible that the genetic information not only fits into the cell but fits into the cell nucleus? Eukaryotes solve this problem by a combination of supercoiling and packaging DNA around the histone family of proteins (described below). Prokaryotes do not contain histones (with a few exceptions). Prokaryotes tend to compress their DNA using nucleoid-associated-proteins (NAPs) and supercoiling, as shown in Figure \(5\). DNA supercoiling refers to the over- or under-winding of a DNA strand and is an expression of the strain on that strand. Supercoiling is important in a number of biological processes, such as compacting DNA and regulating access to the genetic code. DNA supercoiling strongly affects DNA metabolism and possibly gene expression. Additionally, certain enzymes such as topoisomerases can change DNA topology to facilitate functions such as DNA replication or transcription. In a “relaxed” double-helical segment of B-DNA, the two strands twist around the helical axis once every 10.4–10.5 base pairs of sequence. Adding or subtracting twists, as some enzymes can do, impose strain. If a DNA segment under twist strain were closed into a circle by joining its two ends and then allowed to move freely, the circular DNA would contort into a new shape, such as a simple figure-eight, as shown in Figure \(5\). Such a contortion is a supercoil. The noun form “supercoil” is often used in the context of DNA topology. Positively supercoiled (overwound) DNA is transiently generated during DNA replication and transcription, and, if not promptly relaxed, inhibits (regulates) these processes. The simple figure eight is the simplest supercoil and is the shape a circular DNA assumes to accommodate one too many or one too few helical twists. The two lobes of the figure eight will appear rotated either clockwise or counterclockwise with respect to one another, depending on whether the helix is over- or underwound. For each additional helical twist being accommodated, the lobes will show one more rotation about their axis. As a general rule, the DNA of most organisms is negatively supercoiled. Lobal contortions of a circular DNA, such as the rotation of the figure-eight lobes above, are referred to as writhe. The above example illustrates that twisting and writhing are interconvertible. Supercoiling can be represented mathematically by the sum of twist and writhe, The twist is the number of helical turns in the DNA and the writhe is the number of times the double helix crosses over on itself (these are the supercoils). Extra helical twists are positive and lead to positive supercoiling, while subtractive twisting causes negative supercoiling. Many topoisomerase enzymes sense supercoiling and either generate or dissipate it as they change DNA topology. In part, because chromosomes may be very large, segments in the middle may act as if their ends are anchored. As a result, they may be unable to distribute excess twist to the rest of the chromosome or to absorb twist to recover from underwinding—the segments may become supercoiled, in other words. In response to supercoiling, they will assume an amount of writhe, just as if their ends were joined. Supercoiled circular DNA forms two major structures; a plectoneme or a toroid, or a combination of both. A negatively supercoiled DNA molecule will produce either a one-start left-handed helix, the toroid, or a two-start right-handed helix with terminal loops, the plectoneme. Plectonemes are typically more common in nature, and this is the shape most bacterial plasmids will take. For larger molecules, it is common for hybrid structures to form – a loop on a toroid can extend into a plectoneme as shown in Figure \(6\). DNA supercoiling is important for DNA packaging within all cells and seems to also play a role in gene expression. In addition to forming supercoiled structures, circular chromosomes from bacteria have been shown to undergo the processes of catenation and knotting upon the inhibition of topoisomerase enzymes. Catenation is the process by which two circular DNA strands are linked together like chain links, whereas DNA knotting is the interlooping structures occurring within a single circular DNA structure. These are illustrated in Figure \(7\). In vivo, the action of topoisomerase enzymes is critical to keep knots and catenoids from tangling the DNA structure. Mitochondrial and Chloroplast DNA Mitochondrial and Chloroplast DNA are circular suggesting a bacterial origin for both of these organelle structures. Sequence alignments further lend support for the endosymbiotic theory, which proposes that bacteria were engulfed by early eukaryotic organisms and subsequently became symbiotic to their eukaryotic counterpart, rather than being digested. In the cells of eukaryotic organisms, the vast majority of the proteins present in the mitochondria (numbering approximately 1500 different types in mammals) are coded for by nuclear DNA. However, sequencing of the human mitochondrial genome has revealed 16,569 base pairs encoding 13 proteins, as shown in Figure \(8\). Many of the mitochondrially produced proteins are required for electron transport during the production of ATP. Histones and Nucleosomes Within eukaryotic chromosomes, chromatin proteins, known as histones, compact and organize DNA. These compacting structures guide the interactions between DNA and other proteins, helping control which parts of the DNA are transcribed. Histones are highly basic proteins found in eukaryotic cell nuclei that package and order the DNA into structural units called nucleosomes. They are the chief protein components of chromatin, acting as spools around which DNA winds, and playing a role in gene regulation. Without histones, the unwound DNA in chromosomes would be very long (a length-to-width ratio of more than 10 million to 1 in human DNA). For example, each human diploid cell (containing 23 pairs of chromosomes) has about 1.8 meters of DNA wound on the histones, and the diploid cell has about 90 micrometers (0.09 mm) of chromatin. There are five major families of histones, H1/H5, H2A, H2B, H3, and H4. Histones H2A, H2B, H3, and H4 are known as the core histones, while histones H1/H5 are known as the linker histones. The core histones all exist as dimers, which are similar in that they all possess the histone fold domain: three alpha helices linked by two loops (Figure 4.13). It is this helical structure that allows for interaction between distinct dimers, particularly in a head-tail fashion (also called the handshake motif). The resulting four distinct dimers then come together to form one octameric nucleosome core, approximately 63 Angstroms in diameter. Around 146 base pairs (bp) of DNA wrap around this core particle 1.65 times in a left-handed super-helical turn to give a particle of around 100 Angstroms across, called a nucleosome as illustrated in Figure \(9\). The linker histone H1 binds the nucleosome at the entry and exit sites of the DNA, thus locking the DNA into place and allowing the formation of higher order structure (Figure 4.14). The most basic such formation is the 10 nm fiber or beads on a string conformation. This involves the wrapping of DNA around nucleosomes with approximately 50 base pairs of DNA separating each pair of nucleosomes (also referred to as linker DNA). The nucleosome contains over 120 direct protein-DNA interactions and several hundred water-mediated ones. Direct protein – DNA interactions are not spread evenly about the octamer surface but rather located at discrete sites. These are due to the formation of two types of DNA binding sites within the octamer; the α1α1 site, which uses the α1 helix from two adjacent histones, and the L1L2 site formed by the L1 and L2 loops. Salt links and hydrogen bonding between both side-chain basic and hydroxyl groups and main-chain amides with the DNA backbone phosphates form the bulk of interactions with the DNA. This is important, given that the ubiquitous distribution of nucleosomes along genomes requires it to be a non-sequence-specific DNA-binding factor. Although nucleosomes tend to prefer some DNA sequences over others, they are capable of binding practically to any sequence, which is thought to be due to the flexibility in the formation of these water-mediated interactions. In addition, non-polar interactions are made between protein side-chains and the deoxyribose groups, and an arginine side-chain intercalates into the DNA minor groove at all 14 sites where it faces the octamer surface. The spatial distribution and strength of DNA-binding sites about the octamer surface distort the DNA within the nucleosome core. The DNA is non-uniformly bent and also contains twist defects. The twist of free B-form DNA in solution is 10.5 bp per turn. However, the overall twist of nucleosomal DNA is only 10.2 bp per turn, varying from a value of 9.4 to 10.9 bp per turn. The histone tail extensions constitute up to 30% by mass of histones, but are not visible in the crystal structures of nucleosomes due to their high intrinsic flexibility, and have been thought to be largely unstructured (Figure 4.14). The N-terminal tails of histones H3 and H2B pass through a channel formed by the minor grooves of the two DNA strands, protruding from the DNA every 20 bp. The N-terminal tail of histone H4, on the other hand, has a region of highly basic amino acids (16-25), which, in the crystal structure, forms an interaction with the highly acidic surface region of an H2A-H2B dimer of another nucleosome, being potentially relevant for the higher-order structure of nucleosomes. This interaction is thought to occur under physiological conditions also and suggests that acetylation of the H4 tail distorts the higher-order structure of chromatin. Figure \(10\) shows an interactive iCn3D model of the human nucleosome (3afa). One member of each pair of histones is shown in cartoon rendering, while the other member of the pair is shown in the same color but in spacefill rendering. The structure of a human nucleosome (3afa) is shown below (H2A is shown in cyan, H2B in blue, H3 in magenta, and H4 in purple). Each strand of DNA is shown in a different shade of gray. Figure \(10\): Human nucleosome (3afa). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...B2SwQHYDLj4BJ6 The packing of DNA from dsDNA to the metaphase chromosomes is schematically shown in Figure \(11\). The formation of the DNA double helix represents the first-order packaging of the chromosome structure. The formation of nucleosomes represents the second level of packaging for eukaryotic chromosomes. In vitro data suggests that nucleosomes are then arranged into either a solenoid structure which consists of 6 nucleosomes linked together by the Histone H1 linker proteins or a zigzag structure that is similar to the solenoid construct. Both the solenoid and zigzag structures are approximately 30 nm in diameter. The solenoid and zigzag structures reported from in vitro data have not yet been confirmed to occur in vivo. Telomeres At the ends of the linear eukaryotic chromosomes are specialized regions of DNA called telomeres. The main function of these regions is to allow the cell to replicate chromosome ends using the enzyme telomerase, as the enzymes that normally replicate DNA cannot copy the extreme 3′ ends of chromosomes. A cartoon showing telomeres and their extension is shown in Figure \(12\). These specialized chromosome caps also help protect the DNA ends, and stop the DNA repair systems in the cell from treating them as damage to be corrected. In human cells, telomeres contain 300-8000 repeats of a simple TTAGGG sequence. The repetitive TTAGGG sequences in telomeric DNA can form unique higher-order structures called quadruplexes. Figure \(13\) shows an interactive iCn3D model of parallel quadruplexes from human telomeric DNA (1KF1). The structure contains a single DNA strand (5'-AGGGTTAGGGTTAGGGTTAGGG-3') which contains four TTAGGG repeats. Figure \(13\): A buried phenylalanine in low molecular weight protein tyrosyl phosphatase (1xww) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...y5joFHDgWJQsQ6 Rotate the model to see 3 parallel layers of quadruplexes. In each layer, 4 noncontiguous guanine bases interact with a K+ ion. Hover over the guanine bases in one layer and you will find that one layer consists of guanines 4, 10, 16, and 22, which derive from the last G in each of the repeats in the sequence of the oligomer used (5'-AGGGTTAGGGTTAGGGTTAGGG-3'). These quadruplexes certainly serve as recognition and binding sites for telomerase proteins. The guanine-rich telomere sequences which can form quadruplex may also function to stabilize chromosome ends During DNA replication, the double-stranded DNA is unwound and DNA polymerase synthesizes new strands. However, as DNA polymerase moves in a unidirectional manner (from 5’ to 3’), only the leading strand can be replicated continuously. For the complementary lagging strand, DNA replication is discontinuous. In humans, small RNA primers attach to the lagging strand DNA, and the DNA is synthesized in small 5'-3' stretches of about 100-200 nucleotides, which are termed Okazaki fragments. The RNA primers are removed and replaced with DNA and the Okazaki DNA fragments are ligated together. At the end of the lagging strand, it is impossible to attach an RNA primer, meaning that there will be a small amount of DNA lost each time the cell divides. This ‘end replication problem’ has serious consequences for the cell as it means the DNA sequence cannot be replicated correctly, with the loss of genetic information. Hence most telomeres have 3´ overhangs. Bacteria DNA, which is circular, does not have the problem. To prevent this, telomeres are repeated hundreds to thousands of times at the end of the chromosomes. Each time cell division occurs, a small section of telomeric sequences is lost to the end replication problem, thereby protecting the genetic information. At some point, the telomeres become critically short. This decreases leads to cell senescence, where the cell is unable to divide, or apoptotic cell death. Telomeres are the basis for the Hayflick limit, the number of times a cell can divide before reaching senescence. Telomeres can be restored by the enzyme telomerase, which extends the telomere's length. Telomerase activity is found in cells that undergo regular division, such as stem cells and lymphocyte cells of the immune system. The enzyme has two major subunits. One is the catalytic enzyme named telomerase reverse transcriptase (TERT), which is an RNA-dependent DNA polymerase. Almost all DNA polymerases use DNA as a template for replication. Some RNA viruses that use RNA as their genetic information like HIV encode their own reverse transcriptase, which directs the polymerization of a DNA copy of the viral RNA genome as part of its life cycle. )The virus that causes the Covid-19 pandemic is called severe acute respiratory syndrome coronavirus 2 (SARS-CoV-2). It is also an RNA virus but in contrast to HIV it uses and encodes an RNA-dependent RNA polymerase.) The second subunit of telomerase is the telomerase RNA (TR), which contains a template from which the new telomer is made. The enzyme makes many DNA copies from this to create a multitude of DNA repeat in the telomeres. Figure \(14\) shows the structure of the telomerase RNA used to build new telomers. Ovals show proteins that form part of the complex. 'For this discussion, the most important is TERT, telomerase reverse transcriptase, the RNA-dependent DNA polymerase which synthesizes new telomeric DNA from the template sequence of the RNA. Trace the black single-stranded telomeric DNA strand from the 5' end (bottom left) to its 3' end containing the terminal sequence 5'GCTGG3'. Now start tracing the telomerase RNA starting with its 5' end, which is between the CEH and SM binding site in Figure \(14\). It bends sharply near TBE, and continues to the right through the Ku binding site, where it forms stem-loops. It then continues into the template region containing a template sequence (3'CACCGACC5'), from which new telomeric DNA is made (5'GTGGCTGG3' - a slightly different repeat than the human repeat), and an annealing sequence (3'CGACC5'), which is complementary to the 3' end of the existing telomeric DNA (5'GCTGG3'). The RNA continues in the 3' direction through the Est1 bind site region, through the pseudoknot, and ends at the 3' end past the SM binding site. Telomeres can also be extended through the Alternative Lengthening of Telomeres (ALT) pathway. In this case, rather than being extended, telomeres are switched between chromosomes by homologous recombination. As a result of the telomere swap, one set of daughter cells will have shorter telomeres, and the other set will have longer telomeres A downside to telomere extension is the potential for uncontrolled cell division and cancer. Abnormally high telomerase activity has been found in the majority of cancer cells, and non-telomerase tumors often exhibit ALT pathway activation. As well as the potential for losing genetic information, cells with short telomeres are at a higher risk for improper chromosome recombination, which can lead to genetic instability and aneuploidy (an abnormal number of chromosomes). Chromatin Structure During interphase, distinct chromosomes as shown in Figure \(4\) are not observed. Rather each chromosome occupies a spatially limited, roughly elliptical domain which is known as a chromosome territory (CT). Each chromosome territory is comprised of higher-order chromatin units of ~1 Mb each. These units are likely built up from smaller loop domains that contain the solenoid/zigzag structural motifs. On the other hand, 1Mb domains can themselves serve as smaller units in higher-order chromatin structures. With the development of high-throughput biochemical techniques, such as 3C (chromosome conformation capture) and 4C (chromosome conformation capture-on-chip and circular chromosome conformation capture), numerous spatial interactions between neighboring chromatin territories have been described as shown in Figure \(15\). Chromosome territories are known to be arranged radially around the nucleus. This arrangement is both cell and tissue-type specific and is also evolutionarily conserved. The radial organization of chromosome territories was shown to correlate with their gene density and size. In this case, the gene-rich chromosomes occupy interior positions, whereas larger, gene-poor chromosomes, tend to be located around the periphery. Chromosome territories are also dynamic structures, with genes able to relocate from the periphery towards the interior once they have been ‘switched on’. In other cases, genes may move in the opposite direction or simply maintain their position. Figure \(16\) shows nano- to more micro-folding structures of chromosomes in the nucleus. The top left shows the most zoomed-in view where DNA is wound around histone complexes (nucleosomes), which condense to form 10 nm fibers. These condense further into Topologically Associated Domains (TAD) which further separate into Compartments A and B. These then pack into discrete territories. Individual chromosomes occupy their own chromosome territories in the nucleus. This may seem very complex and it is, but it is somewhat analogous to protein folding, which starts with a linear primary sequence and moves into more complicated secondary structures, secondary structure motifs, domains, tertiary structures, and quaternary structures, which display varying degrees of symmetry. Because these different types of folding and compaction of chromosomes are difficult to visualize, we will present several different representations of these nano- to microstructures to help your understanding. Figure \(17\) offers one that shows scaling factors and differential structures of chromosomes and chromatin. The large-scale A compartment is gene-rich and actively transcribed so it best represents euchromatin. In contrast, B compartments are gene-poor and best represent heterochromatin. At the subscale level, boundaries between TADs are transcriptionally rich and can be separated from other TADS by heterochromatin "islands". Another representation of chromatin organization that emphasizes TADs is shown in Figure \(18\). The TAD boundaries in Figure \(18\) show a blue CTCF in-between compartments, with each TAD consisting of many interacting loops. A more detailed view is shown in Figure \(19\). A protein complex containing cohesin (which forms a ring) and CTCF is found at sites where loops of relationally transcribable DNA are extruded through the ring. The extruded loops interact with others to form a cluster of regulatory loops containing genes with similar potential for transcription (activated if they end up in Compartment A or repressed if in Compartment B). Cohesin is a functional complex that forms a ring that traps sister chromatids. During anaphase, the complex is cleaved and the sister chromatids separate. CTCF is a chromatin-binding factor with many associated activities. "Mechanism of chromatin loop formation. TADs contain varying numbers of chromatin loops generated through loop extrusion by CTCF/cohesin complexes. (Right panel) In the presence of NIPBL and MAU2, the cohesin complex loaded onto the DNA. Then, cohesin extrudes chromatin until a pair of convergent CTCF binding sites is reached. (Right panel) The N-terminus of CTCF and convergent positioning of the CTCF-DNA complex stabilizes cohesin binding and stall chromatin extrusion leading to the establishment of higher-order chromatin organization. The intervening DNA between two convergent CTCF sites leads to the formation of a loop domain, which adopts a variety of complex shapes comprised of multiple regulatory loops. The internal structure of the loop domain is likely determined by polymer chromatin-chromatin self-interactions, which may be further stabilized by phase separation. The contacts within the loop domains facilitate the targeting of enhancers to specific genes (104). The black arrow depicts the direction of loop extrusion." Special elements in the DNA called enhancers and silencers of gene transcription have long been known to influence gene transcription. These are sequences that are cis (i.e. on the same molecule of DNA, not trans factors like separate proteins that bind to promoters, for example) and can be quite distant from the proximal promoter sequence which controls the transcription of a target gene. How might they work from such a large distance from the promoter? One obvious answer is the DNA folds in 3D space so the enhancers and silencers are close to each in 3D space. Chromatin loop formation facilitates such promoter and enhancer/silencer interactions. Evidence suggests that specific enhancers and silencers are housed in loops in specific TADs so their effects are limited to a subset of genes. Enhancers and promoters seem to interact only within TADs, which suggests that TADS are a fundamental folding "domain" based on gene regulation. If boundaries between different TADS were removed, then promoters and enhancers in one TAD might affect transcription in other TADS, leading to aberrant gene expression. Chromatin loop formation facilitates interactions between promoter and enhancer/silencer elements. Figure \(20\) shows how enhancers and silencers can be brought close in space to promoters and their genes through TAD formation. TADs appear highly conserved in mammals and comprise most (90%) of the genome. The median size is about 880 Kb. The Boundaries between TADs have CCCTC-binding factor (CTCF) and the structural maintenance of chromosomes (SMC) cohesin complex. In Drosophila, TADs are organized by epigenetic state (methylation, condensation). For example, some TADS are transcriptionally active with epigenetic histone modifications (for example trimethylation of histone H3 lysines 4 and 36 or H3Kme3 and H3K36me3) that activate transcription). Others are transcriptionally repressed (enriched in H3K27me3 and containing Polycomb group (PcG) proteins) and some are more classically representative of heterochromatin. Mechanisms of Separation of Euchromatin and Heterochromatin From a chemistry focus, what are the interactions which stabilize TADs, compartments, and ultimately heterochromatin and euchromatin? It appears that the easiest way to conceptualize their separation in the nucleus is to use the idea of phase separations. Figure \(21\) shows a cartoon representation of the separation of heterochromatin and euchromatin in the zebrafish embryo. The euchromatin is shown as more centrally located and dispersed with red dots indicating transcriptionally active sites. In the late blastula stage, gene expression is dramatically increased and the nucleolus and heterochromatin are not seen. Analyses show that transcription forms regions enriched in RNA, RNA binding proteins, and accordingly transcriptionally active chromatin that are separated from transcriptionally inactive, heterochromatin. Given the dispersion of the DNA required for transcription, the regions enriched in RNA are also depleted in "stainable" DNA. Micrographs showing the depletion of DNA in the actively transcribed regions are shown in Figure \(22\). Note the clear lack of DNA in the white rectangles, which contain high levels of RNA and RNA polymerase II (as a proxy for protein). It appears that the presence of a high concentration of RNA drives "phase" separation of heterochromatin and euchromatin. Hence euchromatin might act as an "oil in water" type microemulsion, with the euchromatin core stabilized by "tethered" RNA acting as an amphiphile. Ribonucleoproteins (RNPs) can be modeled as phase-separated droplets or condensates Figure \(23\) shows how increasing RNA leads to the formation of euchromatin "domains" which stay dispersed in the presence of continuing RNA formation. A final summary cartoon that describes the formation and separation of chromosomal DNA into euchromatin and heterochromatin in Zebra fish is shown in Figure \(24\).

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/08%3A_Nucleotides_and_Nucleic_Acids/8.04%3A_Chromosomes_and_Chromatin.txt

Search Fundamentals of Biochemistry It is difficult to read newspapers and newsmagazines without encountering the CRISPR-Cas9 gene editing system that has the potential to make gene editing routine in disease diagnosis, treatment, and cure, as well as in genetic modification of organisms to improve their quality and quantity for food and natural product production. In this chapter section, we will explore the mechanism of restriction enzymes that made gene cloning possible as well as the CRISPR-Cas gene editing system. Restriction Endonucleases A restriction enzyme, restriction endonuclease, or restrictase is an enzyme that cleaves DNA into fragments at or near specific recognition sites within molecules known as restriction sites. Restriction enzymes are one class of the broader endonuclease group of enzymes. Restriction enzymes are commonly classified into five types, which differ in their structure and whether they cut their DNA substrate at their recognition site, or if the recognition and cleavage sites are separate from one another. To cut DNA, all restriction enzymes make two incisions, once through each sugar-phosphate backbone (i.e. each strand) of the DNA double helix. Here we will focus on the Type II restriction enzymes that are routinely used in molecular biology and biotechnology applications. As with other classes of restriction enzymes, Type II Restriction Enzymes occur exclusively in unicellular microbial life forms––mainly bacteria and archaea (prokaryotes)––and are thought to function primarily to protect these cells from viruses and other infectious DNA molecules. Inside a prokaryote, the restriction enzymes selectively cut up foreign DNA in a process called restriction digestion; meanwhile, host DNA is protected by a modification enzyme (a methyltransferase) that modifies the prokaryotic DNA and blocks cleavage. Together, these two processes form the restriction-modification system. The first Type II Restriction Enzyme discovered was HindII from the bacterium Haemophilus influenzae Rd. The event was described by Hamilton Smith (Figure 7.23) in his Nobel lecture, delivered on 8 December 1978: ‘"In one such experiment we happened to use labeled DNA from phage P22, a bacterial virus I had worked with for several years before coming to Hopkins. To our surprise, we could not recover the foreign DNA from the cells. With Meselson’s recent report in our minds, we immediately suspected that it might be undergoing restriction, and our experience with viscometry told us that this would be a good assay for such an activity. The following day, two viscometers were set up, one containing P22 DNA and the other Haemophilus DNA. Cell extract was added to each and we began quickly taking measurements. As the experiment progressed, we became increasingly excited as the viscosity of the Haemophilus DNA held steady while the P22 DNA viscosity fell. We were confident that we had discovered a new and highly active restriction enzyme. Furthermore, it appeared to require only Mg2+ as a cofactor, suggesting that it would prove to be a simpler enzyme than that from E. coli K or B. After several false starts and many tedious hours with our laborious, but sensitive viscometer assay, Wilcox and I succeeded in obtaining a purified preparation of the restriction enzyme. We next used sucrose gradient centrifugation to show that the purified enzyme selectively degraded duplex, but not single-stranded, P22 DNA to fragments averaging around 100 bp in length, while Haemophilus DNA present in the same reaction mixture was untouched. No free nucleotides were released during the reaction, nor could we detect any nicks in the DNA products. Thus, the enzyme was clearly an endonuclease that produced double-strand breaks and was specific for foreign DNA. Since the final (limit) digestion products of foreign DNA remained large, it seemed to us that cleavage must be site-specific. This proved to be case and we were able to demonstrate it directly by sequencing the termini of the cleavage fragments.’" Restriction enzymes are named according to the taxonomy of the organism in which they were discovered. The first letter of the enzyme refers to the genus of the organism and the second and third to the species. This is followed by letters and/or numbers identifying the isolate. Roman numerals are used to specify different enzymes from the same organism. For example, the enzyme ‘HindIII’ was discovered in Haemophilus influenzae, serotype d, and is distinct from the HindI and HindII endonucleases also present within this bacterium. The DNA-methyltransferases (MTases) that accompany restriction enzymes are named in the same way, and given the prefix ‘M.’. When there is more than one MTase, they are prefixed ‘M1.’, ‘M2.’, etc, if they are separate proteins or ‘M1∼M2.’ when they are joined. Restriction Enzymes that recognize the same DNA sequence, regardless of where they cut, are termed ‘isoschizomers’ (iso = equal; skhizo = split). Isoschizomers that cut the same sequence at different positions are further termed ‘neoschizomers’ (neo = new). Isoschizomers that cut at the same position are frequently, but not always, evolutionarily drifted versions of the same enzyme (e.g. BamHI and OkrAI). Neoschizomers, on the other hand, are often evolutionarily unrelated enzymes (e.g.EcoRII and MvaI). Type II Restriction Enzymes are a conglomeration of many different proteins that, by definition, have the common ability to cleave duplex DNA at a fixed position within, or close to, their recognition sequence. This cleavage generates reproducible DNA fragments, and predictable gel electrophoresis patterns, properties that have made these enzymes invaluable reagents for laboratory DNA manipulation and investigation. Almost all Type II Restriction Enzymes require divalent cations, usually Mg2+, as essential components of their catalytic sites. Ca2+, on the other hand, often acts as an inhibitor of Type II Restriction Enzymes. The recognition sequences of Type II Restriction Enzymes are palindromic, with two possible types of palindromic sequences. The mirror-like palindrome is similar to those found in ordinary text, in which a sequence reads the same forward and backward on a single strand of DNA, as in GTAATG. The inverted repeat palindrome is also a sequence that reads the same forward and backward, but the forward and backward sequences are found in complementary DNA strands (i.e., of double-stranded DNA), as in GTATAC (GTATAC being complementary to CATATG). Inverted repeat palindromes are more common and have greater biological importance than mirror-like palindromes. The position of cleavage within the palindromic sequence can vary depending on the enzyme and can produce either single-stranded overhanging sequences (sticky ends) or blunt-ended DNA products. Examples of staggers and blunt end cuts by restriction enzymes are shown in Table \(8\) below. EcoR1 Sma1 Table \(8\): Staggered and blunt end cut sequences by EcoR1 and Sma1 Methylation can be used by the host to protect its own genome from cleavage. For example, the methylation of the EcoRI recognition sequence by the M.EcoRI methyltransferase (MTase), changes the sequence from GAATTC to GAm6ATTC (m6A = N6-methyladenine). This modification completely protects the sequence from cleavage by EcoRI. Type II Restriction Enzymes initially bind non-specifically with the DNA and proceed to slide down the DNA scanning for recognition sequences as shown in Figure (36\):. Upon binding to the correct palindromic sequence the enzyme associates with the metal cofactor and mediates catalytic cleavage of the DNA using the mechanism of strain distortion and catalysis by approximation. One of the most important questions regarding the catalytic mechanism of a hydrolase is whether hydrolysis involves a covalent intermediate, as is typical for the proteases described previously. This can be decided by analyzing the stereochemical course of the reaction. This was done first for EcoRI, and later for EcoRV. Both enzymes were found to cleave the phosphodiester bond with inversion of the chiral center at the phosphorus, which argues against the formation of a covalent enzyme–DNA intermediate. Thus, it is proposed that cleavage involves the direct nucleophilic attack of the substrate by a water molecule, as shown in Figure (37\) below. Type II restriction enzymes typically form a homodimer when binding with DNA, as shown in the crystal structure of BglII in Figure 7.26B. BglII catalyzes phosphodiester bond cleavage at the DNA backbone through a phosphoryl transfer to water. Studies on the mechanism of restriction enzymes have revealed several general features that seem to be true in almost all cases, although the actual mechanism for each enzyme is most likely some variation of this general mechanism (Figure 7.25). This mechanism requires a base to generate the hydroxide ion from water, which will act as the nucleophile and attack the phosphorus in the phosphodiester bond. Also required is a Lewis acid to stabilize the extra negative charge of the pentacoordinate transition state phosphorus, as well as a general acid or metal ion that stabilizes the leaving group (3’-O). In some Type II Restriction Enzymes, two divalent metal cofactors are required (such as in EcoRV and BamHI), whereas other enzymes only require one divalent metal cofactor (such as in EcoRI and BglII). Structural studies of endonucleases have revealed a similar architecture for the active site with the residues following the weak consensus sequence Glu/Asp-(X)9-20-Glu/Asp/Ser-X-Lys/Glu. BglII's active site is similar to other endonucleases', following the sequence Asp-(X)9-Glu-X-Gln. In its active site, there sits a divalent metal cation, most likely Mg2+, that interacts with Asp-84, Val-94, a phosphoryl oxygen, and three water molecules. One of these water molecules is able to act as a nucleophile because of its proximity to the scissile phosphoryl group (Figure 7.26A). The nucleophilic water molecule is positioned for attack onto the phosphoryl group by a hydrogen bond with the side chain amide oxygen of Gln-95 and its contact with the metal cation. Interaction with the metal cation effectively lowers its pKa, promoting the water's nucleophilicity as shown in Panel A of Figure (38\) below (from Pingoud, A., Wilson, G.G., and Wende, W. (2014) Nuc Acids Res 42(12):7489-7527). During hydrolysis, the divalent cation can stabilize the 3'-O- leaving group and coordinate proton abstraction from one of the coordinated water molecules CRISPR-Cas 9 The CRISPR (clustered regularly interspaced short palindromic repeats) operon was initially discovered as part of the adaptive immune system of bacteria and archaea, which must defend themselves against viruses (bacteriophages) and unwanted plasmids transferred from both bacteria. It would be ideal for bacteria to recognize previous exposure to viruses and their nucleic acids as the basis of their immunological memory system. Given the tendency of viral DNA to integrate into the host genome (which allows later transcription and translations of the viral genes in the process of new virus production), immunological memory could be based on that viral integrated DNA. Without going into detail, viral DNA can be integrated between two direct repeats in the bacterial genome. DNA from different viruses from previous exposures is also incorporated in the same fashion. One site of integration is the CRISPR operon. The DNA of the CRISPR operon contains both protein-coding and noncoding regions which are transcribed and processed to form at least three RNA molecules, as shown in Figure (24\) below. • a coding Cas 9 mRNA this is translated to produce the Cas 9 (CRISPR-associated protein); • a noncoding cr-RNA (CRISPR RNA) • a noncoding tracr-RNA (trans-activating CRISPR RNA) The two mature noncoding RNAs eventually associate to form a binary complex. When using CRISPR-Cas 9 in eukaryotic gene editing applications, the two noncoding RNAs are covalently combined into one large synthetic guide RNA (sg-RNA), described later in this section. The Cas 9 protein is an endonuclease that cleaves both strands of bound target dsDNA in a blunt-end fashion at specific sequences. This occurs after the DNA binds to two arginines (1333 and 1335) in Cas9 through a short (3-5+ bases) recognition protospacer adjacent motif (PAM) located three base pairs from the cleavage site. The DNA must also bind in a complementary and specific fashion to the protein-bound noncoding cr+tracr-RNAs (or a single sg-RNA molecule for gene editing applications). Binding and cleavage of target DNA would render DNA from an invading bacteriophage inactive. Basic research into the bacterial CRISPR system has led to revolutionary and explosive applications of this gene editing system in eukaryotes. The hope is that CRISPR technology will give us a precise and incredibly cheap way to do gene therapy in diseased cells and organisms. Given its role in transforming our ability to edit the genome and potentially cure genetically-based diseases, we will offer a detailed explanation of its mechanism. We have discussed the structure and function of many proteins. Protein enzymes are key to life as they catalyze almost all biological reactions. Most key enzymes are regulated. The activity of Cas 9 must be carefully controlled. Think of the consequences if the enzyme were to cleave promiscuously at off-site targets! This section will help you understand several critical features of this enzyme: 1. How does the enzyme find its correct target site, a 20 nucleotide DNA sequence, and a proximal PAM site, among all the possible alternative sites? Think of how many PAM sequences there must be in the host DNA genome! 2. How can the enzyme be "turned" on when it finds its target site and remain off when free, but more importantly when it is bound off-site? First, we will discuss the apo- form of the enzyme without bound substrate and RNA. Apo- and Holo-Cas 9 This section will focus on the Type II-A Cas9 from Streptococcus pyogenes (SpyCas9 or SpCas9). Cas 9 is an endonuclease that cleaves both strands of DNA 3 base pairs from a DNA motif, NCC/NGG, called PAM. It has two distinct lobes. The nuclease lobe (NUC), amino acids 1-56 and 718-1368, has two different nuclease domains for the two cleavages. The recognition or receptor lobe (REC), amino acids 94-717, interacts with the RNA molecules. There is also an arginine-rich bridge helix (57-93). The enzyme has two catalytic nuclease domains: • HNH-like nuclease domain cleaves the "target" DNA strand, which is complementary to the RNA the confers specificity to the enzyme. The key catalytic residues are His 840 and Asn 854. It also contains a Mg ion; • Ruv-like domain that cleaves the complementary "non-target" strand with key active site residues Asp 10, Glu 762, Asp 986, and His 983. It also contains a bound Mn ion. The two lobes are separated by two linkers, amino acids 712-717, and an arginine-rich bridge (basic helix - BH), amino acids 628-658. The overall structure of the apoenyzme (without bound RNA and DNA,pdb id 4cmp) is shown in Figure (25\) below, which shows the NUC domain (light blue) with the two catalytic domains (HNH and Ruv), the REC domain (orange) and the BH helix (red). A close up view showing the two catalytic sites is shown in Figure (26\) below. Figure \(27\) shows an interactive iCn3D model of Streptococcus pyogenes Cas9 in complex with guide RNA and target DNA (4OO8) (long load time). The Cas9 enzyme is shown as a gray transparent surface with an underlying cartoon rendering. The DNA is shown as colored sticks. The RNA is shown as a cyan cartoon. Figure \(27\): Streptococcus pyogenes Cas9 in complex with guide RNA and target DNA (4OO8). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...RjzJBFVt5qRjS7 (long load time) A comparison of the crystal structure of the apo-Cas 9 and the ternary Cas 9: sgRNA:DNA target strand complex shows a significant conformational change on binding nucleic acids. The structure of the holoenzyme (ternary complex) is shown in Figure (28\) below. The extent of the conformation change between apo- and holo-Cas 9 enzymes can be seen by examining the distance between D435 and E 944/945 in Figure (29\) below. The importance of this change will be described later. Figure (30\) below shows the pathway from the transcription of the relevant CRISPR genes (coding and noncoding) to the assembly of the ternary complex and the blunt end cut of the target DNA strand three nucleotides from the PAM sequence. Figure (31\) below shows an expanded view of the ternary complex. Mechanism of DNA binding and cleavage The above figures do not speak to the mechanism of the binding processes that form the ternary complex. Kinetic and structural studies have been conducted to elucidate the mechanism of binding and cleavage and address the following questions: • which binds first, the RNA or DNA? • What are the consequences of the profound conformational changes on the formation of the ternary complex? The specificity of target DNA binding depends both on enzyme:PAM DNA and enzyme:sgRNA (or tracr- and crRNA) interactions. It should seem improbable that the trinucleotide PAM DNA sequence (NGG in S. pyogenes), which interacts with a pair of arginines (R 1333, R 1335) through H-bonding, as shown in the images above, and other local sites in Cas 9 would provide the sole or even the majority of the binding interactions. Figure (32\) below shows the Args:PAM interaction (pdb code 4un3) Hence it is most likely that RNA binds first. Indeed, it does with the tracRNA implicated in the recruitment of Cas and the crRNA providing specificity for target DNA binding. The resulting Cas9:RNA binary complex could then search the relevant DNA genome. That would include the DNA of the bacteriophage in viral infection or eukaryotic DNA if the CRISPR DNA operon with the genes for Cas 9 and a sg-RNA was transfected into the eukaryotic cell. After RNA binding, the enzyme would change conformation and allow loose DNA binding through Cas 9: PAM interactions. Studies have shown that the apo form can also bind DNA, but it does so loosely and indiscriminately. It dissociates quickly and binding is affected by generic polyanions such as the glycosaminoglycan heparin, which indicates its nonspecific nature. Once bound, both off-target and target DNAs would then be surveyed. If a target DNA contained a PAM sequence, the complex would undergo another conformational change to position the HNH and Ruv nuclease catalytic residues and locally unwind the duplex DNA to make the blunt-end cuts. Cas 9 binding to the PAM site would promote better interaction of the unwound DNA and the bound RNA. If no PAM was present, no catalytically-effective Cas 9:target DNA would form. This prevents off-site cleavage. These allosteric changes and controls are vital to the function of the endonuclease. Here are some findings that support this proposed mechanism: • the conformation of apo Cas 9 is catalytically inactive; • on binding RNA to form a binary complex, Cas 9 undergoes a dramatic conformational change, mostly in the REC lobe. However, on binding DNA in a nonspecific fashion, the conformational changes are much smaller. This suggests that most changes in conformation occur before DNA binding. In a way, RNA acts as an allosteric activator of the enzyme (as well as the major source of binding specificity to target DNA). Conformational changes can be determined directly by comparison of crystal structures or spectral techniques such as fluorescence resonance energy transfer (FRET) between two different attached fluorophores. • Cas 9: RNA interactions lead to ordering of the region of the RNA that interacts with the DNA PAM sequence and adjacent deoxynucleotides (a "seed sequence"), allowing the Cas 9:RNA complex to scan and interact with potential DNA targets with PAM sequences; • Once a PAM site is found, conformational changes lead to unwinding of the dsDNA, which allows heteroduplex formation between the crRNA and the target DNA strand; • since Cas 9 recognizes a variety of DNA target sequences (but of course only a specific PAM sequence), the binding of the target sequence depends on the geometry, not the sequence, of the target DNA; • since binding of off-target DNA to the Cas 9:RNA complex occurs but with very infrequent cleavage, binding and cleavage are very distinct steps; • on specific DNA binding, the HNH catalytic site moves near to the sessile DNA bond site. Crystal structures show that the active site His is not sufficiently close to facilitate cleavage, suggesting that binding of a second metal ion (see below) may be necessary. Molecular dynamics studies show that the HNH domain is "remarkably plastic". Figure (33\) below show an animation that illustrates the relative conformational changes going from the apo Cas 9 to the binary Cas 9:sgRNA complex to the ternary Cas 9: sgRNA: target DNA complex. The NUC catalytic domain is shown in light blue, the REC (receptor or RNA binding domain) in orange, sgRNA in red, and the target DNA in green. Note again that on binding RNA to form a binary complex, Cas 9 undergoes a dramatic conformational change, mostly in the REC lobe. The pdb protein sequences shown were aligned using pdbEfold. A potential abbreviated catalytic mechanism for the Ruv nuclease domain is shown in Figure (34\) below. The red arrows indicate the second set of electron movements. His 983 acts as a general base to abstract a proton from the water making it a more potent nucleophile. An intermediate trigonal bipyramidal phospho-intermediate is formed, which, along with the preceding transition state, is stabilized by the proximal Mg2+ ion (an example of electrostatic or metal ion catalysis). The magnesium is positioned through its interaction with negatively charged carboxyl groups of Asp 10, Glu 762, and Asp 986. A second metal ion might be recruited to the Ruv site to further facilitate the cleavage of the DNA. The HNH catalytic site has a structure (beta-beta-alpha) and conserved His in common with a class of nucleases that require one metal ion. In contrast, the Ruv catalytic site does not have this common secondary structural motif and has a critical histidine, both common features found in endonucleases that use two metal ions. CRISPR and Eukaryotic Gene Editing How could blunt-end cutting of both strands of DNA by Cas 9 lead to the holy grail of specific eukaryotic gene editing with no off-site effects? Cutting the DNA genome seems like a bad idea. It is potentially so bad that a myriad of DNA repair mechanisms has evolved to fix the cut. These include homologous recombination. If corrective DNA is supplied as well as the components of the CRISPR system, a cell could effectively add the corrective DNA after the double-stranded cut and repair a deleterious mutation. Consult a molecular biology textbook for more insight into homologous recombination. Mutations in the PAM sequence prevent Cas9 nuclease activity. Hence the NGG PAM sequence is vital for the interactions and activities described above. This would seem to limit the utility of CRISPR-Cas 9 in eukaryotic gene editing until one realizes that the GG dinucleotide has a 5.2% frequency of occurrence in the human genome, which corresponds to over 160 million occurrences. Even then it might not occur in a desired gene target. Cas 9 nuclease from other bacteria extends the range of activity of the CRISPR/Cas system as they interact with other PAM sequences (NNAGAA and NGGNG for S. thermophilus and NGGNG for N. meningidtis). Likewise, mutations in the S. pyogenes PAM (NGG) have been made as well. A D1135E mutation retains but increases the specificity for the normal NGG PAM site. D1135V, R1335Q, and T1337R mutations alter the optimal PAM recognition site to NGAN or NGNG. CRISPR editing can be easily used to knock out specific genes. In addition, if cells are transfected with a plasmid with many target sequences, the system can be used to edit multiple genes in one experiment. This would be very useful in studies of diseases linked to multiple genes. Since the cost of CRISPR reagents (plasmids, RNAs) is so inexpensive, and the specificity of editing is so high, the great excitement about CRISPR use for gene editing in human disease and for modification of plant and fungal genomes is warranted. Other systems have been developed to specifically bind to a target DNA sequence and then cleave it. They typically contain a protein that binds to a specific DNA target and an associated endonuclease that cleaves within the target DNA site. Typical prokaryotic restriction enzymes bind to and cut at a specific nucleotide sequence (for example Eco R1 cleaves at G/AATTC palindromic sequences) to form sticky ends. The protein itself binds to this DNA recognition site. Other examples are based on the structure of known transcription factors. Libraries of genetically engineered proteins with Zn finger DNA binding domains (designed for specific DNA target sequences) fused to endonucleases have been created for this purpose. Other examples are proteins called TALENs (transcription activator-like effector nucleases). These are fusion proteins containing a TAL effector DNA-binding domain and a nuclease. In each of these cases, a 3D-folded protein is the specific target DNA recognition molecule. Think how much easier it is to make in effect a 1D-DNA recognition element, a simple linear RNA sequence, which would adopt the correct 3D structure on the binding of its complementary target. One major problem in the use of CRISPR for gene editing must be solved: how to get the CRISPR components in the correct cells in an organism. In effect, it's the same problem faced by small drug designers only the components are much larger. Ex vivo applications, when diseased cells are removed from the body, repaired by CRISPR, and then reinjected, are likely to have more success. In these cases, electroporation would allow the uptake of Cas 9 and the sg-RNA. In vivo therapy has included the use of adeno-associated viruses in which genes for Cas 9 and sg RNA could be encapsulated. This technique, used for other gene delivery systems, has the advantage of being tolerated immunologically. However, this system allows for continual gene expression which is undesirable for gene editing. After an initial "fix" of a mutant gene, continued expression of the CRISPR-Cas 9 genes would increase the chances for off-target cutting. A more recent approach is to deliver the mRNA in artificial lipid nanoparticles that can be taken into cells. Once free and translated into protein and sg RNA inside the cell, gene editing has a chance to occur before the RNA and protein are degraded.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/08%3A_Nucleotides_and_Nucleic_Acids/8.06%3A_Enzymes_for_Genetic_modifications.txt

• 9.1: DNA Isolation, Sequencing, and Synthesis • 9.2: Bioinformatics • 9.3: Cloning and Recombinant Expression • 9.4: DNA Microarrays A DNA microarray (also commonly known as DNA chip or biochip) is a collection of microscopic DNA spots attached to a solid surface. Scientists use DNA microarrays to measure the expression levels of large numbers of genes simultaneously or to genotype multiple regions of a genome. Each DNA spot contains picomoles (10−12 moles) of a specific DNA sequence, known as probes (or reporters or oligos). • 9.5: In Situ Hybridization In situ hybridization (ISH) is a type of hybridization that uses a labeled complementary DNA, RNA or modified nucleic acids strand (i.e., probe) to localize a specific DNA or RNA sequence in a portion or section of tissue (in situ) or if the tissue is small enough (e.g., plant seeds, Drosophila embryos), in the entire tissue (whole mount ISH), in cells, and in circulating tumor cells (CTCs). This is distinct from immunohistochemistry, which usually localizes proteins in tissue sections. • 9.6: References 09: Investigating DNA Search Fundamentals of Biochemistry Genomic and complementary DNA The ability to sequence the DNA of an organism has revolutionized our understanding of biology and evolution. DNA can be isolated directly from living, dead, and even extinct species and the "primary" sequence of A, G, C, and T bases in the molecule determine. We can read (sequence), write (synthesize), and edit (mutate) DNA at will. Before we explore how to purify, sequence (read) and synthesize (write) DNA, it's important to differentiate two different types of DNAs, genomic and complementary DNA (cDNA), which is made by reverse transcribing messenger RNA into DNA. Since mRNA has no nucleotides encoded by introns, c-DNA provides just the coding sequences for protein. Genomic deoxyribonucleic acid (gDNA) is chromosomal DNA, which does not include the extra-chromosomal DNA found in the mitochondria of eukaryotes or plasmids in bacteria (plasmids will be discussed in more detail in section 5.3 during the discussion of gene cloning and expression). Most organisms have the same genomic DNA in every cell (one exception is the genomic DNA for antibodies in B cells and T cells receptors in T-cells that have been altered as the cells become more terminally differentiated. It is also important to remember that only certain genes are active (expressed) in each cell. The subset of expressed genes is specific to a given differentiated cell and allows for the expression of specific cell functions. Liver cells, for example, don't express the gene for the protein opsin, which is expressed in retinal cells and is required for vision. The genome of an organism (encoded by the genomic DNA) is the (biological) information of heredity that is passed from one generation of organism to the next. That genome is transcribed to produce various RNAs, which are necessary for the function of the organism. Precursor mRNA (pre-mRNA) is transcribed by RNA polymerase II in the nucleus. pre-mRNA is then processed by splicing to remove introns, leaving the exons in the mature messenger RNA (mRNA). Additional processing includes the addition of a 5' cap and a poly(A) tail to the pre-mRNA. The mature mRNA may then be transported to the cytosol and translated by the ribosome into a protein. Other types of RNA include ribosomal RNA (rRNA) and transfer RNA (tRNA). These types are transcribed by RNA polymerase II and RNA polymerase III, respectively, and are essential for protein synthesis. However, 5s rRNA is the only rRNA that is transcribed by RNA Polymerase III. cDNA is derived from mRNA, so it contains only exons but no introns. Figure \(1\) shows the flow of information stored in eukaryotic DNA and its eventual expression in mRNA. Red indicates coding exons, which are separated by gray introns. At the beginning and end of a gene sequence (encoded by 3 exons in the figure below) are 5' and 3' untranslated regions (UTRs) which are also transcribed and are represented in the mature mRNA. Also, there are potential regulatory sequences, which are not transcribed (yellow) on both sides of the transcribable part of the gene. The 5'-end promoter is where transcription factors and RNA polymerase assemble before the start of transcriptions. In addition, there are regulatory enhancers and silencers more distal from the gene sequences. An open reading frame (ORF) is a region of the DNA that can be decoded into a mRNA and doesn't have a stop signal (codon) in it that would prematurely terminate transcription. In contrast, complementary DNA (cDNA) is synthesized from a single-stranded RNA (e.g., messenger RNA (mRNA) or microRNA) template in a reaction catalyzed by the enzyme reverse transcriptase. Reverse transcriptase is an enzyme found in retroviruses such as HIV that have RNA as their core genetic material. Upon entering the host cell the RNA is reverse-transcribed to produce a copy of cDNA that can then integrate into the host's genomic DNA. In biotechnology, reverse transcriptase is often used to create cDNA from the mRNA expressed in specific cells or tissues. In this way, the eukaryotic genes can be cloned without any introns housed in the structure. This is especially useful if the goal is to express the protein in a prokaryotic (bacterial) host. Recall that bacterial DNA does not contain any intron sequences within its chromosomal DNA. Thus, if you are using a prokaryotic system to express eukaryotic proteins, you must use cDNA, as the prokaryotic system will not be able to remove intron sequences following gene transcription. The term cDNA is also used, typically in a bioinformatics context, to refer to an mRNA transcript's sequence expressed as DNA bases (GCAT) rather than RNA bases (GCAU). The gene organization of prokaryotes is different in that they don't have introns. In addition, some genes for a common pathway for example are continuous in the DNA. These stretches of DNA are called operons. Transcription from an operon produces a polycistronic RNA transcript. The words cis and are used in chemistry to describe R groups on the same size (cis) or opposite sides (trans) of a double bond. In DNA, cis-elements are in a single DNA section while trans-elements usually refer to proteins (away from the gene) binding to the DNA. Hence the term polycistronic for bacterial operons (with multiple genes sequentially arranged in the DNA sequence). Figure \(2\) shows the organization of prokaryotic gene structure. DNA Extraction/Purification The first isolation of DNA was done in 1869 by Friedrich Miescher. Now purification kits are available from multiple manufacturers. DNA can be isolated from whole tissue or cell cultures. Let's consider just DNA extraction from cells grown in the lab. Cells are first collected by centrifugation and then treated with detergents like sodium dodecyl sulfate to lyse the cell membranes. Proteases and DNAase-free RNAase can be added to digest proteins and RNA, respectively. Methods involving phenol/chloroform extractions: In older methods, a mixture of phenol and chloroform or phenol/chloroform/isoamyl alcohol is used to extract DNA from the solution. Students who have performed liquid/liquid extractions in chemistry lab courses should recognize that this will form a biphasic mixture with water. Nonpolar substances like lipids and cellular debris will partition into the nonpolar phase (chloroform/phenol) or in the interface between them (as suspended insoluble material). Chloroform is very dense as it contains a chlorine atom). Phenol is somewhat soluble in water (8 g/100 g water) but very soluble in chloroform. In mixing during the extraction, the dissolved phenol alters the properties of water sufficiently enough to push the delicate equilibrium of proteins from the native to the denatured states, which aggregate and precipitate. On settling, DNA will remain in the aqueous phase. The use of chloroform/phenol in DNA extractions has a potential problem. Phenol (hydroxylated benzene) can lose one electron from the oxygen atom forming a free radical, which can be stabilized by resonance with pi electrons in the aromatic ring. Free radicals can damage DNA, so most new methods of purification do not use phenol/chloroform extractions. Most methods involve precipitating the extracted DNA at some point in the purification process using cold ice-cold ethanol or isopropanol. DNA is to a first approximation a long polyanion so it would be very difficult to purify "naked" DNA from solution since the extensive negative charges on the DNA would prevent aggregate and precipitate formation. This is not a problem if the ionic strength of the medium is sufficiently high so bound positively charged counter-ions can shield the negative charges from each other, allowing precipitation. Methods involving adsorption chromatography using silica gel: Nucleic acids bind or adsorb to a solid phase (silica or other) depending on the pH and the salt concentration of the buffer. If small amounts are needed (such as for isolating recombinant plasmid from bacteria), small spin columns are employed. This method relies on the fact that nucleic acid will bind to the solid phase of silica gel under certain conditions and then be released when those conditions are altered. These features are illustrated in Figure \(3\). Figure \(3\): DNA-Silica gel interactions. Image by Squidonius The solution containing DNA is applied to a small spin column containing silica gel and placed in a mini-centrifuge. On spinning, the nucleic acid will bind to the silica gel membrane as the solution passes through. After multiple "spin" washes to remove non-specific cellular components from the column, the DNA is eluted with a low salt elution buffer (or simply water). Unlike RNA which degrades very quickly, DNA is quite stable and can be stored for long periods at -20oC. Many student readers have used silica gel chromatography or silica gel thin-layer chromatography to separate and analyze organic mixtures. These techniques are usually performed in a mixture of organic solvents (like hexane/ethyl acetate for example). The use of silica gel to purify DNA from an aqueous solution might seem strange so we will briefly explore how DNA binds. In silica gel, each silicon atom is tetrahedrally bonded (sp3 hybridization) to four oxygen atoms, with each oxygen atom covalently linked to two silicon atoms. At the surface of the particle, the oxygen atoms are capped with H atoms, so the entire surface contains a "sea" of OHs, and therefore hydrogen bond donors and acceptors. At lower pHs, some could ionize to form O- ions. If salt concentrations are high enough, the percentage of O- ions increases, since they are stabilized by the cations in the salt (shifting the equilibrium to the ionized state). DNA, a long negatively charged anion, can bind to the silica surface using two types of noncovalent interaction. It can form hydrogen bonds with the silica gel surface OHs. In addition, it can interact with the surface through ion-ion interactions mediated by bridging cations (like Na+ from the high-concentration salt solution) as illustrated in the figure above. The binding solutions used in the spin column adsorption steps have high concentrations of chaotropic salts that disrupt water structure and hydrogen bonding. The salts also denature proteins and in effect dehydrate the DNA. Some chaotropic salts used include sodium iodide, sodium perchlorate, guanidinium thiocyanate, and guanidinium chloride. The sodium or guanidinium acts as bridging cations, allowing the adsorption of the negative DNA to negative charges on the silica gel surface. Sodium acetate and Tris-HCl are included to buffer pH from 6-7. Now it becomes easy to understand how pure water or low salt concentration solutions elute the bound DNA after extensive column washing since pure water or low salt solutions would strip the bound intermediary cations from the silica column. After isolation, the DNA is dissolved in a slightly alkaline buffer, usually in a Tris-EDTA buffer, or in ultra-pure water. EDTA binds divalent cations like Ca2+, which activate nucleases. Modifications made to these standard techniques are often done if the tissue being used is difficult to break down, if contaminants persist in the lysis solution that inhibit further reactions, or if the sample is extremely minimal, as is often the case in forensic investigations. In addition, different commercial kits will be tailored for the isolation of larger genomic DNA or smaller plasmid DNA. The purity of a DNA preparation is usually determined by measuring the absorbance of the solution at 230, 260 (peak absorbance nucleic acids), and 280 nm (peak absorbance proteins), often using an instrument that requires a tiny droplet of solution. Figure (4\) below shows the relative absorbance spectra of proteins and nucleic acids. Figure (4\) relative absorbance spectra of proteins and nucleic acids. Brianna Bibel. The Bumbling Chemist https://thebumblingbiochemist.com/36...purity-ratios/ The A260/A280 gives a measure of protein contamination, with a value around 1.8 indicating "pure" DNA. The A260/A230 gives information on protein and other solution contamination. A pure solution of DNA that has an A260 = 1.0 has a concentration of 50 μg/mL = 50 ng/μL. When the temperature of a dsDNA solution is increased, in some range of temperature the A260 increases by about 37%. This is called the hyperchromic effect which occurs when the bases in DNA unstack on denaturation of dsDNA to single-stranded DNA as the intrastrand hydrogen bonds break. Figure (5\) below shows a graph of A260 vs temperature. The midpoint of the cooperative change in the absorbance signifies that the population of DNA molecules is 50% denatured. Given the cooperative of unfolding, it is less likely that the population consists of individual molecules that are 50% denatured (i.e. a single molecule being 50% double-stranded and 50% single-stranded). Accordingly, a lower concentration (33 μg/mL) of fully single-stranded DNA gives an A260 = 1. DNA Sequencing Techniques DNA sequencing is the process of determining the nucleic acid sequence – the order of nucleotides in DNA. It includes any method or technology that is used to determine the order of the four bases: adenine, guanine, cytosine, and thymine. The advent of rapid DNA sequencing methods has greatly accelerated biological and medical research and discovery. Knowledge of DNA sequences has become indispensable for basic biological research and in numerous applied fields such as medical diagnosis, biotechnology, forensic biology, virology, and biological systematics. Comparing healthy and mutated DNA sequences can diagnose different diseases including various cancers, characterize antibody repertoire, and can be used to guide patient treatment. Having a quick way to sequence DNA allows for faster and more individualized medical care to be administered, and for more organisms to be identified and cataloged. The rapid speed of sequencing attained with modern DNA sequencing technology has been instrumental in the sequencing of complete DNA sequences, or genomes, of numerous types and species of life, including the human genome and other complete DNA sequences of many animal, plant, and microbial species. The first DNA sequences were obtained in the early 1970s by academic researchers using laborious methods based on two-dimensional chromatography. Following the development of fluorescence-based sequencing methods with a DNA sequencer, DNA sequencing has become easier and orders of magnitude faster. The canonical structure of DNA has four bases: thymine (T), adenine (A), cytosine (C), and guanine (G). DNA sequencing is the determination of the physical order of these bases in a molecule of DNA. However, DNA bases are often modified by epigenetic processes to control gene expression. Thus, many other modified bases may be present in a DNA molecule than the standard four bases. For example, in some viruses (specifically, bacteriophages), cytosine may be replaced by hydroxymethyl- or hydroxymethylglucose cytosine. In eukaryotic DNA, variant bases with methyl groups or phosphosulfate may be found as shown in Figure (6\) below. Depending on the sequencing technique, a particular modification, e.g., the 5mC (5 -methylcytosine) common in humans, may or may not be detected. Early DNA sequencing methods The first method for determining DNA sequences involved a location-specific primer extension strategy established by Ray Wu at Cornell University in 1970. DNA polymerase catalysis and specific nucleotide labeling, both of which figure prominently in current sequencing schemes, were used to sequence the cohesive ends of lambda phage DNA. Between 1970 and 1973, Wu, R Padmanabhan, and colleagues demonstrated that this method can be employed to determine any DNA sequence using synthetic location-specific primers. Frederick Sanger then adopted this primer-extension strategy to develop more rapid DNA sequencing methods at the MRC Centre, Cambridge, UK, and published a method for "DNA sequencing with chain-terminating inhibitors" in 1977. Walter Gilbert and Allan Maxam at Harvard also developed sequencing methods, including one for "DNA sequencing by chemical degradation". In 1973, Gilbert and Maxam reported the sequence of 24 base pairs using a method known as wandering-spot analysis. Advancements in sequencing were aided by the concurrent development of recombinant DNA technology, allowing DNA samples to be isolated from sources other than viruses. Maxam-Gilbert sequencing requires radioactive labeling at one 5' end of the DNA and purification of the DNA fragment to be sequenced. Chemical treatment then generates breaks at a small proportion of one or two of the four nucleotide bases in each of the four reactions (G, A+G, C, C+T). The concentration of the modifying chemicals is controlled to introduce on average one modification per DNA molecule. Thus a series of labeled fragments is generated, from the radiolabeled end to the first "cut" site in each molecule. The fragments in the four reactions are electrophoresed side by side in denaturing acrylamide gels for size separation. To visualize the fragments, the gel is exposed to X-ray film for autoradiography, yielding a series of dark bands each corresponding to a radiolabeled DNA fragment, from which the sequence may be inferred. The technical aspects of Maxam-Gilbert sequencing caused it to go out of favor once the Sanger sequencing method had been well established, as described below. Sanger Sequencing Method The chain-termination method developed by Frederick Sanger and coworkers in 1977 soon became the method of choice, owing to its relative ease and reliability. When invented, the chain-terminator method used fewer toxic chemicals and lower amounts of radioactivity than the Maxam-Gilbert method. Because of its comparative ease, the Sanger method was soon automated and was the method used in the first generation of DNA sequencers. The classical chain-termination method requires a single-stranded DNA template, a DNA primer, a DNA polymerase, normal deoxynucleotide triphosphates (dNTPs), and modified di-deoxynucleotide triphosphates (ddNTPs), the latter of which terminate DNA strand elongation. These chain-terminating nucleotides lack a 3'-OH group required for the formation of a phosphodiester bond between two nucleotides, causing DNA polymerase to cease the extension of DNA when a modified ddNTP is incorporated. The ddNTPs may be radioactively or fluorescently labeled for detection in automated sequencing machines. The DNA sample is divided into four separate sequencing reactions, containing all four of the standard deoxynucleotides (dATP, dGTP, dCTP, and dTTP) and the DNA polymerase. To each reaction is added only one of the four dideoxynucleotides (ddATP, ddGTP, ddCTP, or ddTTP), while the other added nucleotides are ordinary ones as shown in (7\) below. The dideoxynucleotide concentration should be approximately 100-fold lower than that of the corresponding deoxynucleotide (e.g. 0.005mM ddTTP : 0.5mM dTTP) to allow enough fragments to be produced while still transcribing the complete sequence. In total, four separate reactions are needed in this process to test all four ddNTPs. This is illustrated in Figure (8\) below. Following rounds of template DNA extension from the bound primer, the resulting DNA fragments are heat denatured and separated by size using gel electrophoresis. This technique was frequently performed using a denaturing polyacrylamide-urea gel with each of the four reactions run in one of four individual lanes (lanes A, T, G, C). The DNA bands may then be visualized by autoradiography or UV light and the DNA sequence can be directly read off the X-ray film or gel image, as shown in Figure (9\) below. Figure (9\): Traditional Sanger Sequencing Gel. Sequence visualized by autoradiography. Each lane contains a single reaction that has all four regular nucleotides and a small amount of one of the dideoxynucleotides (ddNTPs). Over time, the ddNTPs will be incorporated at each position containing that specific nucleotide. The gel can then be read from the bottom to the top, as the smallest fragments (those fragments terminated the closest to the primer at the 5'-end) will run the farthest distance in the gel. The sequence of this fragment is 5'-TACGAGATATATGGCGTTAATACGATATATTGGAACTTCTATTGC-3'. Image by John Schmidt Automation of the Sanger sequencing method was made possible when the shift from radioactively tagged nucleotides to fluorescently tagged nucleotides was made. Within the automated sequencers, capillary gel electrophoresis is performed rather than separating the samples using gel electrophoresis. The output from capillary electrophoresis are fluorescent peak trace chromatograms, as shown in Figure (10\) below. Automated DNA-sequencing instruments (DNA sequencers) can sequence up to 384 DNA samples in a single batch. Batch runs may occur up to 24 times a day greatly enhancing the speed with which samples may be sequenced and analyzed. Common challenges of DNA sequencing with the Sanger method include poor quality in the first 15-40 bases of the sequence due to primer binding and deteriorating quality of sequencing traces after 400-500 bases. Figure (10\): Side by Side Comparison of Gel Electrophoresis and Capillary Electrophoresis. Lefthand Diagram shows the traditional autoradiogram of Sanger sequencing samples. The Righthand Diagram shows the same reactions using fluorescently tagged ddNTPs separated by capillary electrophoresis. The chromatogram output is shown on the far right. Image by Abizar Sanger sequencing is the method that prevailed from the 1980s until ~2005. Over that period, great advances were made in the technique, such as fluorescent labeling, capillary electrophoresis, and general automation. These developments allowed much more efficient sequencing, leading to lower costs. The Sanger method, in mass production form, is the technology that produced the first human genome in 2001, ushering in the age of genomics. Microfluidic Sanger Sequencing Microfluidic Sanger sequencing is a lab-on-a-chip application for DNA sequencing, in which the Sanger sequencing steps (thermal cycling, sample purification, and capillary electrophoresis) are integrated on a wafer-scale chip using nanoliter-scale sample volumes (Figure (11\)). This technology generates long and accurate sequence reads while obviating many of the significant shortcomings of the conventional Sanger method (e.g. high consumption of expensive reagents, reliance on expensive equipment, personnel-intensive manipulations, etc.) by integrating and automating the Sanger sequencing steps. Next-generation sequencing (NGS) Next-generation sequencing (NGS), also known as high-throughput sequencing, is the catch-all term used to describe several different modern sequencing technologies. These technologies allow for the sequencing of DNA and RNA much more quickly and cheaply than the previously used Sanger sequencing, and as such revolutionized the study of genomics and molecular biology. We present information from specific companies that have developed these new technologies without endorsements. Such technologies include: Illumina Sequencing - In NGS, vast numbers of short reads are sequenced in a single stroke using the lab-on-a-chip technology described above. To do this, the input sample must be cleaved into short sections. In Illumina sequencing, 100-150bp reads are used. Somewhat longer fragments are ligated to generic adaptors and annealed to a slide using the adaptors. PCR is carried out to amplify each read, creating a spot with many copies of the same read. They are then separated into single-stranded DNA to be sequenced as shown in Figure (12\) below. Roche 454-Sequencing is similar to the Illumina process but can sequence much longer reads. Like Illumina, it does this by sequencing multiple reads at once by reading optical signals as bases are added. As in Illumina, the DNA or RNA is fragmented into shorter reads, in this case, up to 1kb (1,000bp). Generic adaptors are added to the ends and these are annealed to beads, one DNA fragment per bead. The fragments are then amplified by PCR using adaptor-specific primers. Each bead is then placed in a single well of a slide with each well containing a single bead, covered in many PCR copies of a single sequence. The wells also contain DNA polymerase and sequencing buffers (Figure (13\). Newer technologies such as the Ion Torrent Technology detect sequence data using electrical signals on a semiconductor chip, rather than optically reading dye-labeled nucleotides. This is possible as the addition of a dNTP to the DNA polymer causes the release of an H+ ion (Figure (14\)). As in other kinds of NGS, the input DNA or RNA is fragmented, this time ~200bp. Adaptors are added and one molecule is placed onto a bead. The molecules are amplified on the bead by emulsion PCR. Each bead is placed into a single well of a slide. This is illustrated in Figure (14\) below. Nanopore Sequencing In this technique, flow cells are constructed which contain nanopores in a nonlipid membrane that is resistant to current flow. When a voltage is applied across a membrane separating two salt solutions, the current through each nanopore channel can be detected by a sensor chip. When a larger molecule moves through the pore, a disruption in basal current occurs. Computer algorithms have been developed that detect base-specific changes in the current as the base (even chemical-modified ones) moves through the membrane. The sequence is then decoded in real-time. Single-stranded DNA or RNA can be driven through the pore by a transmembrane potential in a process similar to electrophoresis. The enzyme DNA helicase, a motor protein, can be attached to the outer part of the pore protein. This enzyme binds single-stranded DNA and moves along the DNA in a process that requires ATP. If the helicase is attached to the pore protein, the single DNA would then move, allowing control of its movement through the pore. The nanopores are made of real membrane proteins (which we discuss in Chapter 11). One example is α-hemolysin, a heptamer that has an inner pore diameter of 1 nm. When embedded in real cells, it can allow the flow of K+ (diameter around 250 pm = 0.25 nm) and other ions across the cell membrane, changing the osmotic balance and lysing the cell. The pore size of the proteins used in nanopore sequencing allows single-stranded DNA to flow through it. An interactive iCn3D model of protein used for nanopore sequencing, Curli transport lipoprotein CsgG (4uv3), is shown in the membrane bilayer in Figure \(15\). Figure \(15\): Curli transport lipoprotein CsgG (4uv3) for DNA nanopore sequencing. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...LC5K3vSjYjtHN7 Nanopore technologies have enabled the production of small hand-held DNA sequencing devices that can be plugged into the USB drive on a laptop computer and utilized in the field under real-time collection conditions. Future modifications might replace protein pores with synthetic solid-state nanopores. For instance, the membrane might be made of graphene with pores of a specific size made in it. Figure \(16\) shows an animation of a single-stranded DNA moving through a protein pore (blue) assisted by a motor protein (magenta) Single Molecule, Real-Time (SMRT) sequencing In this technique, either RNA or DNA is converted to dsDNA. Deoxynucleotide "adapters" are added to connect the 5'-end of strand 1 to the 3'-end of strand 2 and another adapter to connect the 3'-end of strand 1 to the 5'-end of strand 2 resulting in a "circular" ss-DNA molecule. This single molecule is then drawn into a nanophotonic nanowell made in a thin metal film deposited on glass. The dimensions of each well allow only a single circular ss-DNA molecule. Hundred of different circular ss-DNA molecules entering individual wells are shown in Figure \(17\). The blue stretches represent the adapters. The wells are approximately 100 nm in diameter. DNA polymerase and dNTPs can be added to the nanowells, which contained a single molecule of immobilized circular ssDNA. The DNA is immobilized by its biotinylated or attachment to magnetic beads which interact with streptavidin-coated wells. When confined to the wells the apparent concentration of reactants for the polymerization can be quite high, allowing robust DNA polymerase activity. DNA sequencing using real-time fluorescence monitoring can be done in a massively parallel fashion. The fluorophore is connected to the terminal phosphate of the dNTP. When a phosphodiester bond is made by the DNA polymerase, the fluorophore is released as a leaving group, and the result is a natural, unmodified DNA that continues to grow. The YouTube video below shows the entire process of a single molecule, real-time sequencing. The four main advantages of Next Generation Sequencing (NGS) over classical sequencing are described below. Sample size NGS is significantly cheaper, quicker, needs significantly less DNA, and is more accurate and reliable than Sanger sequencing. Let us look at this more closely. For Sanger sequencing, a large amount of template DNA is needed for each read. Several strands of template DNA are needed for each base being sequenced (i.e. for a 100bp sequence you'd need many hundreds of copies, for a 1000 bp sequence you'd need many thousands of copies), as a strand that terminates on each base is needed to construct a full sequence. In NGS, a sequence can be obtained from a single strand. In both kinds of sequencing multiple staggered copies are taken for contig construction and sequence validation. Speed NGS is quicker than Sanger sequencing in two ways. Firstly, the chemical reaction may be combined with the signal detection in some versions of NGS, whereas in Sanger sequencing these are two separate processes. Secondly and more significantly, only one read (maximum ~1kb) can be taken at a time in Sanger sequencing, whereas NGS is massively parallel, allowing 300Gb of DNA to be read on a single run on a single chip. Cost The reduced time, people power, and reagents in NGS mean that the costs are much lower. The first human genome sequence cost in the region of \$2.7 billion in 2003. Using modern Sanger sequencing methods, aided by data from the known sequence, a full human genome still cost \$300,000 in 2006. Sequencing a human genome with NGS today costs roughly \$1,000. Accuracy Repeats are intrinsic to NGS, as each read is amplified before sequencing, and because it relies on many short overlapping reads, each section of DNA or RNA is sequenced multiple times. Also, because it is so much quicker and cheaper, it is possible to do more repeats than with Sanger sequencing. More repeats mean greater coverage, which leads to a more accurate and reliable sequence, even if individual reads are less accurate for NGS. The nanopore and single molecule, real-time (SMRT) sequencing were recently employed to complete the sequence for the full human genome (2022). Previous genomic sequences were missing regions with highly repetitive sequences at the centromeres and telomeres. The "Telomere-to-Telomere (T2T) Consortium performed the analysis which added 200 megabases of new sequence information that was missing from the previous best sequence. DNA Synthesis Techniques DNA synthesis is the natural or artificial creation of deoxyribonucleic acid (DNA) molecules. The term DNA synthesis can refer to DNA replication (which will be covered in more detail in Chapter XX), polymerase chain reaction (PCR), or gene synthesis (physically creating artificial gene sequences). Polymerase Chain Reaction (PCR) The Polymerase chain reaction (PCR) refers to a technique employed widely in the basic and biomedical sciences. PCR is a laboratory technique utilized to amplify specific segments of DNA for a wide range of laboratory and/or clinical applications. Building on the work of Panet and Khorana’s successful amplification of DNA in-vitro, Kary Mullis and coworkers developed PCR in the early 1980s, having been met with a Nobel prize only a decade later. Allowing for more than the billion-fold amplification of specific target regions, it has become instrumental in many applications including the cloning of genes, the diagnosis of infectious diseases, and the screening of prenatal infants for deleterious genetic abnormalities. Fundamentals The main components of PCR are a template, primers, free nucleotide bases, and the DNA polymerase enzyme. The DNA template contains the specific region that you wish to amplify, such as the DNA extracted from a piece of hair for example. Primers, or oligonucleotides, are short strands of single-stranded DNA complementary to the 3' end of each target region. Both a forward and a reverse primer are required, one for each complementary strand of DNA. DNA polymerase is the enzyme that carries out DNA replication. Thermostable analogs of DNA polymerase I, such as Taq polymerase, which was originally found in a bacterium that grows in hot springs, are a common choice due to their resistance to the heating and cooling cycles necessary for PCR. PCR takes advantage of the complementary base pairing, double-stranded nature, and melting temperature of DNA molecules. This process involves cycling through 3 sequential rounds of temperature-dependent reactions: DNA melting (denaturation), annealing, and enzyme-driven DNA replication (elongation). Denaturation begins by heating the reaction to about 95oC, disrupting the hydrogen bonds that hold the two strands of template DNA together. Next, the reaction is reduced to around 50 to 65oC, depending on the physicochemical variables of the primers, enabling the annealing of complementary base pairs. The primers, which are added to the solution in excess, bind to the beginning of the 3' end of each template strand and prevent re-hybridization of the template strand with itself. Lastly, enzyme-driven DNA replication, or elongation, begins by setting the reaction temperature to the amount which optimizes the activity of DNA polymerase, which is around 75 to 80oC. At this point, DNA polymerase, which needs double-stranded DNA to begin replication, synthesizes a new DNA strand by assembling free nucleotides in solution in the 3' to 5' direction to produce 2 full sets of complementary strands. The newly synthesized DNA is now identical to the template strand and will be used as such in the progressive PCR cycles. The steps in PCR are animated in the video below. Figure (17\) below illustrate the step involved in PCR amplification of target DNA. Figure (18\) shows a video animation of a PCR reaction. Figure (18\): Video animation of a PCR reaction. Given that previously synthesized DNA strands serve as templates, the amplification of DNA using PCR increases at an exponential rate, where the copies of DNA double at the end of each replication step. The exponential replication of the target DNA eventually plateaus around 30 to 40 cycles mainly due to reagent limitation, but can also be due to inhibitors of the polymerase reaction found in the sample, self-annealing of the accumulating product, and accumulation of pyrophosphate molecules. Real-Time PCR At its advent, PCR technology was limited to qualitative and or semi-quantitative analysis due to limitations on the ability to quantitate nucleic acids. At that time, to verify if the target gene was amplified successfully, the DNA product was separated by size via agarose gel electrophoresis. Ethidium bromide, a molecule that fluoresces when bound to dsDNA, could give a rough estimate of DNA amount by roughly comparing the brightness of separated bands, but was not sensitive enough for rigorous quantitative analysis. Improvements in fluorophore development and instrumentation led to thermocyclers that no longer required measurement of only end-product DNA. This process, known as real-time PCR, or quantitative PCR (qPCR), has allowed for the detection of dsDNA during amplification. qPCR thermocyclers are equipped with the ability to excite fluorophores at specific wavelengths, detect their emission with a photodetector, and record the values. The sensitive collection of numerical values during amplification has strongly enhanced quantitative analytical power. There are two main types of fluorophores used in qPCR: those that bind specifically to a given target sequence and those that do not. The sensitivity of fluorophores has been an important aspect of qPCR development. One of the most effective and widely used non-specific markers, SYBR Green, after binding to the minor groove of dsDNA, exhibits a 1000-fold increase in fluorescence compared to being free in solution (Video 5.1). However, if even more specificity is desired, a sequence-specific oligonucleotide, or hybridization probe, can be added, which binds to the target gene at some point in front of the primer (after the 3' end). These hybridization probes contain a reporter molecule at the 5' end and a quencher molecule at the 3' end. The quencher molecule effectively inhibits the reporter from fluorescing while the probe is intact. However, upon contact with DNA polymerase I, the hybridization probe is cleaved, allowing for the fluorescence of the dye (Video 5.1). Reverse-Transcription PCR Since its advent, PCR technology has been creatively expanded upon, and reverse-transcription PCR (RT-PCR) is one of the most important advances. Real-time PCR is frequently confused with reverse-transcription PCR, but they are separate techniques. In RT-PCR, the DNA amplified is derived from mRNA by using reverse-transcriptase enzymes, to produce a cDNA copy of the gene. Using primer sequences for genes of interest, traditional PCR methods can be used with the cDNA to study the expression of genes qualitatively. Currently, reverse-transcription PCR is commonly used with real-time PCR, which allows one to quantitatively measure the relative change in gene expression across different samples. Figure (19\) shows a video animation video showing the use of reverse transcription Polymerase Chain Reaction (RT-PCR) in COVID-19 testing. Figure (19\): Video animation of the Reverse Transcription Polymerase Chain Reaction (RT-PCR) Issues of Concern One disadvantage of PCR technology is that it is extremely sensitive. Trace amounts of RNA or DNA contamination in the sample can produce extremely misleading results. Another disadvantage is that the primers designed for PCR require sequence data, and therefore can only be used to identify the presence or absence of a known pathogen or gene. Another limitation is that sometimes the primers used for PCR can anneal non-specifically to similar sequences, but not identical, to the target gene. Another potential issue of using PCR is the possibility of primer dimer (PD) formation. PD is a potential by-product and consists of primer molecules that have hybridized with each other due to the strings of complementary bases in the primers. The DNA polymerase amplifies the PD, leading to competition for PCR reagents that could be used to amplify the target sequences. Clinical Significance PCR amplification is an indispensable tool with various applications within medicine. Often, it is used to test for the presence of specific alleles, such as in the case of prospective parents screening for genetic carriers, but it can also be used to diagnose the presence of disease directly and for mutations in the developing embryo. For example, the first time PCR was used in this way was for the diagnosis of sickle cell anemia through the detection of a single gene mutation. Additionally, PCR has greatly revolutionized the diagnostic potential for infectious diseases, as it can be used to rapidly determine the identity of microbes that were traditionally unable to be cultured, or that required weeks for growth. Pathogens routinely detected using PCR include Mycobacterium tuberculosis, human immunodeficiency virus, herpes simplex virus, syphilis, and countless other pathogens. Moreover, qPCR is not only used for testing the qualitative presence of microbes but also to quantify bacterial, fungal, and viral loads. The sensitivity of diagnostic tools for mutations to oncogenes and tumor suppression genes has been improved at least 10,000-fold due to PCR, allowing for earlier diagnosis of cancers like leukemia. PCR has also enabled more nuanced and individualized therapies for cancer patients. Additionally, PCR can be used for the tissue typing done that is vital to organ implantation and has even been proposed as a replacement for antibody-based tests for blood type. PCR also has clinical applications in the field of prenatal testing for various genetic diseases and/or clinical pathologies. Samples are obtained either via amniocentesis or chorionic villus sampling. In forensic medicine, short pieces of repeating, highly polymorphic DNA, coined short tandem repeats (STRs), are amplified and used to compare specific variations within genes to differentiate individuals.[9] Primers specific for the loci of these STRs are used and amplified using PCR. Various loci contain STRs in the human genome, and the statistical power of this technique is enhanced by checking multiple sites. Gene Synthesis Artificial gene synthesis, sometimes known as DNA printing is a method in synthetic biology that is used to create artificial genes in the laboratory. Based on solid-phase DNA synthesis, it differs from molecular cloning and polymerase chain reaction (PCR) in that it does not have to begin with preexisting DNA sequences. Therefore, it is possible to make a completely synthetic double-stranded DNA molecule with no apparent limits on either nucleotide sequence or size. The method has been used to generate functional bacterial or yeast chromosomes containing approximately one million base pairs. Creating novel nucleobase pairs in addition to the two base pairs in nature could greatly expand the genetic code. Synthesis of the first complete gene, a yeast tRNA, was demonstrated by Har Gobind Khorana and coworkers in 1972. Synthesis of the first peptide- and protein-coding genes was performed in the laboratories of Herbert Boyer and Alexander Markham, respectively. Commercial gene synthesis services are now available. Approaches are most often based on a combination of organic chemistry and molecular biology techniques and entire genes may be synthesized "de novo", without the need for template DNA. Gene synthesis is an important tool in many fields of recombinant DNA technology including heterologous gene expression, vaccine development, gene therapy, and molecular engineering. The synthesis of nucleic acid sequences can be more economical than classical cloning and mutagenesis procedures. It is also a powerful and flexible engineering tool for creating and designing new DNA sequences and protein functions. Gene optimization While the ability to make increasingly long stretches of DNA efficiently and at lower prices is a technological driver of this field, increasing attention is being focused on improving the design of genes for specific purposes. Early in the genome sequencing era, gene synthesis was used as an (expensive) source of cDNAs that were predicted by genomic or partial cDNA information but were difficult to clone. As higher-quality sources of sequence-verified cloned cDNA have become available, this practice has become less urgent. Producing large amounts of protein from gene sequences can sometimes prove difficult. Many of the most interesting proteins are normally regulated to be expressed in very low amounts in wild-type cells. Redesigning these genes offers a means to improve gene expression in many cases. Rewriting the open reading frame is possible because of the degeneracy of the genetic code. Thus it is possible to change up to about a third of the nucleotides in an open reading frame and still produce the same protein. The available number of alternate designs possible for a given protein is astronomical. For a typical protein sequence of 300 amino acids, there are over 10150 codon combinations that will encode an identical protein. Codon optimization, or replacing rarely used codons with more common codons sometimes has dramatic effects. Further optimizations such as removing RNA secondary structures can also be included. At least in the case of E. coli, protein expression is maximized by predominantly using codons corresponding to tRNA that retain amino acid charging during starvation. Computer programs are used to optimize this task. A well-optimized gene can improve protein expression 2 to 10-fold, and in some cases, more than 100-fold improvements have been reported. Because of the large number of nucleotide changes made to the original DNA sequence, the only practical way to create the newly designed genes is to use gene synthesis. Oligonucleotide synthesis Oligonucleotides are chemically synthesized using building blocks called nucleoside phosphoramidites. These can be normal or modified nucleosides that have protecting groups to prevent their amines, hydroxyl groups, and phosphate groups from interacting incorrectly. One phosphoramidite is added at a time, the 5' hydroxyl group is deprotected and a new base is added, and the process is repeated. The chain grows in the 3' to 5' direction, which is backward relative to DNA biosynthesis in vivo. In the end, all the protecting groups are removed. The solid phase DNA synthesis reaction is shown in Figure (20\) below. Nevertheless, being a chemical process, several incorrect interactions occur leading to some defective products. The longer the oligonucleotide sequence that is being synthesized, the more defects there are, thus this process is only practical for producing short sequences of nucleotides. The current practical limit is about 200 bp (base pairs) for an oligonucleotide with sufficient quality to be used directly for a biological application. HPLC can be used to isolate products with the proper sequence. Meanwhile, a large number of oligos can be synthesized in parallel on gene chips. For optimal performance in subsequent gene synthesis procedures, they should be prepared individually and on larger scales. DNA synthesis and synthetic biology The significant drop in the cost of gene synthesis in recent years due to increasing competition of companies providing this service has led to the ability to produce entire bacterial plasmids that have never existed in nature. The field of synthetic biology utilizes the technology to produce synthetic biological circuits, which are stretches of DNA manipulated to change gene expression within cells and cause the cell to produce a desired product. The ability to synthetically produce DNA will enable the development of environmental, medical, and commercially relevant products. For example, in 2015, Novartis, in collaboration with the Synthetic Genomics Vaccines inc. and the US Biomedical Advanced Research and Development Authority, announced that they had effectively created a synthetic DNA influenza vaccine. New synthetic DNA vaccines hold promise to provide an alternative to current egg-produced conventional vaccines that can be plagued by low efficacy. DNA vaccines can avoid many issues associated with egg-based vaccine production by generating viral proteins within host cells. To create a DNA vaccine, an antigen-encoding gene is cloned into a non-replicative expression plasmid, which is delivered to the host by traditional vaccination routes. Host cells that take up the plasmid express the vaccine antigen which can be presented to immune cells via the major histocompatibility complex (MHC) pathways. CD4+ T helper cell activation following MHC class II presentation of secreted DNA vaccine protein is critical for the production of antigen-specific antibodies as shown in Figure (21\) below. After two decades of research, DNA vaccine technology is gaining maturity—several veterinary DNA vaccines are currently licensed for West Nile virus and melanoma, and significantly, the first commercial DNA vaccine against H5N1 in chickens has recently been conditionally approved by the USDA. In addition, ongoing large animal trials of DNA vaccines against other diseases such as HIV, hepatitis, and Zika virus offer valuable insights that can be applied to influenza DNA vaccine design. Promising approaches have arisen from the numerous studies evaluating different DNA vaccine formulations and delivery systems, but a strategy that consistently elicits protection against influenza in large animal models has not yet emerged. Successful plasmid delivery and the use of appropriate adjuvants remain key challenges that need to be addressed before influenza DNA vaccines become effective for human use. RNA Vaccines Of course, amazing progress has been made in the creation of RNA vaccines as demonstrated by the Moderna and Pfizer RNA vaccines to the spike protein of the SARS-CoV2 virus. Models show that just in the US  through 2022, Covid vaccines saved the US \$1.15 trillion and 3 million lives.  Just in the first year, the vaccines are estimated to have saved upwards of 20 million lives worldwide, an accomplishment worthy of every prize in the world!  Many more could have been saved in the developing world if the vaccine was more widely available. As we learned previously, RNA is much more labile to hydrolysis than DNA since RNA has a 2' OH group. Methods to stabilize RNA were required before an RNA vaccine could become a reality. More importantly, dsRNA is a danger signal that a viral infection may be present. dsRNA, a pathogen-associated molecular pattern (PAMP), binds to the Toll-like Receptor 3 (TLR3) and initiates an inflammatory response that would eliminate the RNA before it could be decoded to form a protein sequence that could elicit an antibody response, a requirement for a vaccine. Katalin Karikó and Drew Weissman found that modifying the uracil bases to pseudouracil (Ψ) prevents the PAMP response and allows RNA to last long enough to create the protein sequence (i.e. it increases the stability of the RNA to hydrolysis). Pseudouracil (Ψ) is found in structural RNAs (transfer, ribosomal, small nuclear, and small nucleolar), and in fact is the most common modification found in RNA. It even is metabolized by a naturally occurring pathway back to uracil. Figure (22\) below shows the structures of uracil and the N1-methyl derivative of pseudouracil attached to ribose in an RNA and their base pairing to adenine. The N1-methylpseudouracil base still bases pairs with an adenine base.  Hence the RNA structure and its functional ability to be decoded into a protein sequence is not affected by the modification.  In short, mRNAs modified to contain pseudouracil have a much greater translational capacity as well as stability.  In addition, methylation of uracil decreases the immunogenicity of RNAs that contain it. Figure (23\) below shows a cartoon version of the modified mRNA (a) Covid-19 and its encapsulation into a lipid nanoparticle (b) for an mRNA vaccine. Panel (a) shows the generalized structure of the mRNA vaccine for the S gene which encodes the surface spike protein of the virus. Panel b shows the lipid nanoparticle that encapsulates and protects the mRNA vaccine. The lipids include proprietary mixtures of phospholipids, cholesterol, cationic (ionizable) lipids, and polyethylene glycol (PEGylated). Without the basic research of Karikó and Weissman (and many others), the world would not have had the Covid-19 vaccine in time to save so many lives. They were awarded the Nobel Prize in Medicine in 2023 for their work. Biosynthesis of pseudouridine The biosynthesis of pseudouridine, the most common modification of cellular RNA, is done after DNA transcription (i.e. post-transcriptionally) using the enzyme pseudouridine synthases (PUS), which is found in all kingdoms in life. The reaction involves • cleavage of the C-N-glycosidic bond of uridine in RNA • rotation of the cleaved uracil to align C5 of uracil and C1′ of the ribose • formation of the C1′-C5 carbon–carbon bond. These processes are illustrated in Figure (24\) below. Figure (24\): Post-transcriptional modification of uridine to pseudouridine.  Czudnochowski N. et al. The mechanism of pseudouridine synthases from a covalent complex with RNA, and alternate specificity for U2605 versus U2604 between close homologs. Nucleic Acids Res. 2014 Feb;42(3):2037-48. doi: 10.1093/nar/gkt1050. Epub 2013 Nov 7. PMID: 24214967; PMCID: PMC3919597. Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0/)

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/09%3A_Investigating_DNA/9.01%3A_DNA_Isolation_Sequencing_and_Synthesis.txt

Search Fundamentals of Biochemistry Introduction An unprecedented revolution has been observed in science with recent technological advances, which have provided a large amount of “omic” data. The crescent generation and availability of this information available in public databases were, and still are, a challenge for professionals from different areas. However, what is the challenge? In biology, the main challenge is to make sense of the enormous amount of structural data and sequences that have been generated at multiple levels of biological systems. Still, in bioinformatics, the development of tools is necessary (statistical and computational) capable of assisting in understanding the mechanisms underlying biological questions in the study. Besides, if we consider the complexity of science, this is a highly reductionist view. The era of a “new biology” emerges accompanied by the birth/development of other sciences, such as bioinformatics and computational biology, which have an integrated interface with molecular biology. Although considered recently, bioinformatics and genomics have evolved interdependently and promoted a historical impact on the available knowledge. Bioinformatics is a hybrid science that links biological data with techniques for information storage, distribution, and analysis to support multiple areas of scientific research including biomedicine. It uses molecular biology and genetics, computer science, mathematics, and statistics to address data-intensive, large-scale biological problems are addressed from a computational point of view. Bioinformatics is fed by high-throughput data-generating experiments, including genomic sequence determinations and measurements of gene expression patterns. Database projects curate and annotate the data and then distribute it via the World Wide Web. Mining these data leads to scientific discoveries and in the end to the identification of new clinical applications. A bioinformatics solution usually involves the following steps: • Collect statistics from biological data • Build a computational model • Solve a computational modeling problem • Test and evaluate a computational algorithm It also addresses the following aspects: • Types of biological information and databases • Sequence analysis and molecular modeling • Genomic analysis • Systems biology In the field of medicine in particular, many important applications for bioinformatics have been discovered. For example, it is used to identify correlations between gene sequences and diseases, to predict protein structures from amino acid sequences, to aid in the design of novel drugs, and to tailor treatments to individual patients based on their DNA sequences (pharmacogenomics). Using bioinformatics, we can now conduct global analyses of all the available data to uncover common principles that apply across many systems and highlight novel features. Many bioinformatic tools are available online in the fields of genomics, comparative genomics, proteomics, drug discovery, cancer research, phylogenetics, forensic sciences, biodefense, nutrigenomics, gene expressions, protein structure/function, etc. Instead of endorsing any given tool, and constantly curating web addresses, which often change, we offer a series of links that readers can explore to find the tools appropriate to their needs. 9.03: Cloning and Recombinant Expression Search Fundamentals of Biochemistry To clone a gene from an organism and express it in either a prokaryotic or eukaryotic cells, DNA from a target source must be isolated, purified, amplified, analyzed and sequenced as described in previous sections. Cloning In general, cloning means the creation of a perfect replica. Typically, the word is used to describe the creation of a genetically identical copy. In biology, the re-creation of a whole organism is referred to as “reproductive cloning.” Long before attempts were made to clone an entire organism, researchers learned how to copy short stretches of DNA—a process that is referred to as molecular cloning. Molecular cloning allows for the creation of multiple copies of genes, the expression of genes, and study the of specific genes. To get the DNA fragment into a bacterial cell in a form that will be copied or expressed, the fragment is first inserted into a cloning vector. Cloning vector A Cloning vector small piece of DNA that can be stably maintained in an organism, and into which a foreign DNA fragment can be inserted for cloning purposes. The cloning vector may be DNA taken from a virus, the cell of a higher organism, or the plasmid of a bacterium. The vector contains features that allow for the convenient insertion or removal of a DNA fragment to or from the vector, for example by treating the vector and the foreign DNA with a restriction enzyme that cuts the DNA. DNA fragments thus generated contain either blunt ends or overhangs known as sticky ends, and vector DNA and foreign DNA with compatible ends can then be joined together by molecular ligation. After a DNA fragment has been cloned into a cloning vector, it may be further subcloned into another vector designed for more specific use. There are many types of cloning vectors, but the most commonly used ones are genetically engineered plasmids. Cloning is generally first performed using Escherichia coli, and cloning vectors in E. coli include plasmids, bacteriophages (such as phage λ), cosmids, and bacterial artificial chromosomes (BACs). Some DNA, however, cannot be stably maintained in E. coli, for example very large DNA fragments. For these studies, other organisms such as yeast may be used. Cloning vectors in yeast include yeast artificial chromosomes (YACs). The common bacterial cloning plasmid, pRB322, is shown in Figure \(1\). All commonly used cloning vectors in molecular biology have key features necessary for their function, such as a suitable cloning site with restriction enzymes and a selectable marker. Others may have additional features specific to their use. For reasons of ease and convenience, cloning is often performed using E. coli. Thus, the cloning vectors used often have elements necessary for their propagation and maintenance in E. coli, such as a functional origin of replication (ori). The ColE1 origin of replication is found in many plasmids. Some vectors also include elements that allow them to be maintained in another organism in addition to E. coli, and these vectors are called shuttle vectors. Cloning site All cloning vectors have features that allow a gene to be conveniently inserted into the vector or removed from it. This may be a multiple cloning site (MCS) or polylinker, which contains many unique restriction sites. The restriction sites in the MCS are first cleaved by restriction enzymes, then a PCR-amplified target gene also digested with the same enzymes is ligated into the vectors using DNA ligase. The target DNA sequence can be inserted into the vector in a specific direction if so desired. The restriction sites may be further used for sub-cloning into another vector if necessary. Other cloning vectors may use topoisomerase instead of ligase and cloning may be done more rapidly without the need for restriction digest of the vector or insert. In this TOPO cloning method, a linearized vector is activated by attaching topoisomerase I to its ends, and this "TOPO-activated" vector may then accept a PCR product by ligating both the 5' ends of the PCR product, releasing the topoisomerase and forming a circular vector in the process. Another method of cloning without the use of a DNA digest and ligase is by DNA recombination, for example as used in the Gateway cloning system. The gene, once cloned into the cloning vector (called entry clone in this method), may be conveniently introduced into a variety of expression vectors by recombination. Restriction Enzymes Restriction enzymes (also called restriction endonucleases) recognize specific DNA sequences and predictably cut them; they are naturally produced by bacteria as a defense mechanism against foreign DNA. As the name implies, restriction endonucleases (or restriction enzymes) are “restricted” in their ability to cut or digest DNA. The restriction that is useful to biochemists is usually a palindromic DNA sequence. Palindromic sequences are the same sequence forwards and backward. Some examples of palindromes: RACE CAR, CIVIC, A MAN A PLAN A CANAL PANAMA. DNA has two complementary strands. Therefore, the reverse complement of one strand is identical to the other. Like with a palindromic word, the DNA palindromic sequence reads the same forward and backward. In most cases, the sequence reads the same forward on one strand and backward on the complementary strand. Restriction enzymes often cut DNA into a staggered pattern. When a staggered cut is made in a sequence, the overhangs are complementary as shown in Figure \(2\). Figure \(2\): Restriction Enzyme Recognition Sequences. In this (a) six-nucleotide restriction enzyme recognition site, notice that the sequence of six nucleotides reads the same in the 5′ to 3′ direction on one strand as it does in the 5′ to 3′ direction on the complementary strand. This is known as a palindrome. (b) The restriction enzyme makes breaks in the DNA strands and (c) the cut in the DNA results in “sticky ends”. Another piece of DNA cut on either end by the same restriction enzyme could attach to these sticky ends and be inserted into the gap made by this cut. http://opentextbc.ca/biology/wp-cont...e_10_01_04.jpg Molecular biologists also tend to use these special molecular scissors that recognize palindromes of 6 or 8. By using 6-cutters or 8-cutters, the sequences occur rarely, but often enough, to be of utility. Figure \(3\) the sequence for HindII cuts. Figure \(3\): Sequence of HindIII stick end cuts. Figure \(4\) shows restriction enzyme cuts that leave sticky or blunt end. Figure \(4\): Restriction Enzymes. Restriction enzymes recognize palindromic sequences in DNA and hydrolyze covalent phosphodiester bonds of the DNA to leave either “sticky/cohesive” ends or “blunt” ends. This distinction in cutting is important because an EcoRI sticky end can be used to match up a piece of DNA cut with the same enzyme to glue or ligate them back together. While endonucleases cut DNA, ligases join them back together. DNA digested with EcoRI can be ligated back together with another piece of DNA digested with EcoRI, but not to a piece digested with SmaI. Another blunt cutter is EcoRV with a recognition sequence of GAT | ATC. Selectable marker A selectable marker is carried by the vector to allow the selection of positively transformed cells. Antibiotic resistance is often used as a marker , an example being the beta-lactamase gene, which confers resistance to the penicillin group of beta-lactam antibiotics like ampicillin. Some vectors contain two selectable markers, for example, the plasmid pACYC177 has both ampicillin and kanamycin resistance genes. Shuttle vectors that are designed to be maintained in two different organisms may also require two selectable markers, although some selectable markers such as resistance to zeocin and hygromycin B are effective in different cell types. Auxotrophic selection markers that allow an auxotrophic organism to grow in a minimal growth medium may also be used; examples of these are LEU2 and URA3 which are used with their corresponding auxotrophic strains of yeast. Another kind of selectable marker allows for the positive selection of plasmid with cloned genes. This may involve the use of a gene lethal to the host cells, such as barnase, Ccda, and the parD/parE toxins. This typically works by disrupting or removing the lethal gene during the cloning process, and unsuccessful clones where the lethalstill remains intact would kill the host cells, therefore only successful clones are selected. Reporter genes Reporter genes are used in some cloning vectors to facilitate the screening of successful clones by using features of these genes that allow successful clone clones easily identified. Such features present in cloning vectors may be the lacZα fragment for α complementation in the blue-white selection, and/or marker gene or reporter genes in frame with and flanking the MCS to facilitate the production of fusion proteins. Examples of fusion partners that may be used for screening are the green fluorescent protein (GFP) and luciferase. Figure \(5\) shows such a construct with GFP. Elements for expression If the expression of the targeted gene is desired, then a cloning vector also needs to contain suitable elements for the expression of the cloned target gene, including a promoter and ribosomal binding site (RBS). The target DNA may be inserted into a site that is under the control of a particular promoter necessary for the expression of the target gene in the chosen host. Where the promoter is present, the expression of the gene is preferably tightly controlled and inducible so that proteins are only produced when required. Some commonly used promoters are the T7 and lac promoters. The presence of a promoter is necessary when screening techniques such as blue-white selection are used. Cloning vectors without promoter and RBS for the cloned DNA sequence are sometimes used, for example when cloning genes whose products are toxic to E. coli cells. Promoter and RBS for the cloned DNA sequence are also unnecessary when first making a genomic or cDNA library of clones since the cloned genes are normally subcloned into a more appropriate expression vector if their expression is required. Types of cloning vectors A large number of cloning vectors are available, and choosing the right vector may depend on a number of factors, such as the size of the insert, copy number, and cloning method. Large DNA inserts may not be stably maintained in a general cloning vector, especially for those with a high copy number, therefore cloning large fragments may require a more specialized cloning vector. Plasmids Plasmids are autonomously replicating circular extra-chromosomal DNA. They are the standard cloning vectors and the ones most commonly used. Most general plasmids may be used to clone DNA inserts to 15 kb in size. Many plasmids have high copy numbers, for example, pUC19 has a copy number of 500-700 copies per cell, and a high copy number is useful as it produces a greater yield of recombinant plasmid for subsequent manipulation. However low-copy-number plasmids may be preferably used in certain circumstances, for example, when the protein from the cloned gene is toxic to the cells. Bacteriophage The bacteriophages most commonly used for cloning are the lambda (λ) phage and M13 phage. There is an upper limit on the amount of DNA that can be packed into a phage (a maximum of 53 kb). The average lambda phage genome is roughly 48.5 kb. Therefore to allow foreign DNA to be inserted into phage DNA, phage cloning vectors may need to have some of their non-essential genes deleted to make room for the foreign DNA. The phage sequence and cartoon structure are shown in Figure \(6\). There is also a lower size limit for DNA that can be packed into a phage. This property can be used for selection - vectors without inserts may be too small, therefore only vectors with inserts may be selected for propagation. Figure \(6\): Lambda Phage. (A) Schematic representation of the circular genome of the lambda phage (B) Diagram of the Lambda Phage infectious particle and (C) Electron micrograph of the related bacteriophage, vibriophage VvAWI. The bar denotes 50 nm in length. Images A and C are modified from Nigro, O, Culley, A., and Steward, G.F. (2012) Standards in Genomic Science 6(3):415-26, and image B is from Jack Potte Cosmids Cosmids are plasmids that incorporate a segment of bacteriophage λ DNA that has the cohesive end sites (cos) which contain elements required for packaging DNA into λ particles. It is normally used to clone large DNA fragments between 28 and 45 Kb. Bacterial artificial chromosome Insert size of up to 350 kb can be cloned in a bacterial artificial chromosome (BAC). BACs are maintained in E. coli with a copy number of only 1 per cell. BACs have often been used to sequence the genome of organisms in genome projects, including the Human Genome Project. A short piece of the organism's DNA is amplified as an insert in BACs, and hen sequenced. Finally, the sequenced parts are rearranged in silico, resulting in the genomic sequence of the organism. BACs have largely been replaced in this capacity with faster and less laborious sequencing methods like whole genome shotgun sequencing and now more recently next-gen sequencing. Yeast artificial chromosome Yeast artificial chromosomes are used as vectors to clone DNA fragments of more than 1 megabase (1Mb = 1000kb = 1,000,000 bases) in size. They are useful in cloning larger DNA fragments as required in mapping genomes such as in the human genome project. It contains a telomeric sequence, an autonomously replicating sequence( features required to replicate linear chromosomes in yeast cells). These vectors also contain suitable restriction sites to clone foreign DNA as well as genes to be used as selectable markers. Human artificial chromosome Human artificial chromosomes may be potentially useful as gene transfer vectors for gene delivery into human cells, and a tool for expression studies and determining human chromosome function. It can carry very large DNA fragments (there is no upper limit on size for practical purposes), therefore it does not have the problem of limited cloning capacity of other vectors, and it also avoids possible insertional mutagenesis caused by integration into host chromosomes by a viral vector. Animal and plant viral vectors that infect plant and animal cells have also been manipulated to introduce foreign genes into plant and animal cells. The natural ability of viruses to adsorb to cells, introduce their DNA and replicate has made them ideal vehicles to transfer foreign DNA into eukaryotic cells in culture. A vector based on Simian virus 40 (SV40) was used in first the cloning experiment involving mammalian Severall vectors based on viruses like Adenoviruses and Papilloma virus have been used to clone genes in mammals. At present, retroviral vectors are popular for cloning genes in mammalian cells. In the case of plant transformation, viruses including the Cauliflower Mosaic Virus, Tobcco Mosaic Virus, and Gemini Viruses have been used with limited success. Summary of DNA Cloning Figure \(7\) provides a summary of the basic cloning methods most widely used in biochemistry laboratories. Foreign DNA is isolated or amplified using PCR to obtain enough material for the cloning procedure. The DNA is purified and cut with restriction enzymes, and then mixed with a vector that has been cut with the same restriction enzymes. The DNA can then be stitched back together with DNA ligase. The DNA can then be transformed into a host system, often bacteria, to grow large quantities of the plasmid containing the cloned DNA. Restriction fragment patterning and DNA sequencing can be used to validate the cloned material. For a Video Tutorial on DNA Cloning visit HHMI - BioInteractive. Plasmids with foreign DNA inserted into them are called recombinant DNA molecules because they contain new combinations of genetic material. Proteins that are produced from recombinant DNA molecules are called recombinant proteins. Not all recombinant plasmids are capable of expressing genes. Plasmids may also be engineered to express proteins only when stimulated by certain environmental factors, so that scientists can control the expression of the recombinant proteins. Reproductive Cloning Reproductive cloning is a method used to make a clone or an identical copy of an entire multicellular organism. Most multicellular organisms undergo reproduction by sexual means, which involves the contribution of DNA from two individuals (parents), making it impossible to generate an identical copy or a clone of either parent. Recent advances in biotechnology have made it possible to reproductively clone mammals in the laboratory. Natural sexual reproduction involves the union, during fertilization, of a sperm and an egg. Each of these gametes is haploid, meaning they contain one set of chromosomes in their nuclei. The resulting cell, or zygote, is then diploid and contains two sets of chromosomes. This cell divides mitotically to produce a multicellular organism. However, the union of just any two cells cannot produce a viable zygote; there are components in the cytoplasm of the egg cell that are essential for the early development of the embryo during its first few cell divisions. Without these provisions, there would be no subsequent development. Therefore, to produce a new individual, both a diploid genetic complement and an egg cytoplasm are required. The approach to producing an artificially cloned individual is to take the egg cell of one individual and to remove the haploid nucleus. Then a diploid nucleus from a body cell of a second individual, the donor, is put into the egg cell. The egg is then stimulated to divide so that development proceeds. This sounds simple, but it takes many attempts before each of the steps is completed successfully. The first cloned agricultural animal was Dolly, a sheep who was born in 1996 (see Figure \(8\) below). The success rate of reproductive cloning at the time was very low. Dolly lived for six years and died of a lung tumor. There was speculation that because the cell DNA that gave rise to Dolly came from an older individual, the age of the DNA may have affected her life expectancy. Since Dolly, several species of animals (such as horses, bulls, and goats) have been successfully cloned. There have been attempts at producing cloned human embryos as sources of embryonic stem cells. In the procedure, the DNA from an adult human is introduced into a human egg cell, which is then stimulated to divide. The technology is similar to the technology that was used to produce Dolly, but the embryo is never implanted into a surrogate mother. The cells produced are called embryonic stem cells because they can develop into many different kinds of cells, such as muscle or nerve cells. The stem cells could be used to research and ultimately provide therapeutic applications, such as replacing damaged tissues. The benefit of cloning in this instance is that the cells used to regenerate new tissues would be a perfect match to the donor of the original DNA. For example, a leukemia patient would not require a sibling with a tissue match for a bone-marrow transplant. To create Dolly, the nucleus was removed from a donor egg cell. The enucleated egg was placed next to the other cell, then they were shocked to fuse. They were shocked again to start division. The cells were allowed to divide for several days until an early embryonic stage was reached, before being implanted in a surrogate mother. Why was Dolly a Finn-Dorset and not a Scottish Blackface sheep? Because even though the original cell came from a Scottish Blackface sheep and the surrogate mother was a Scottish Blackface, the DNA came from a Finn-Dorset. Genetic Engineering Genetic engineering uses recombinant DNA technology to modify an organism’s DNA to achieve desirable traits. The addition of foreign DNA in the form of recombinant DNA vectors that are generated by molecular cloning is the most common method of genetic engineering. An organism that receives the recombinant DNA is called a genetically modified organism (GMO). If the foreign DNA that is introduced comes from a different species, the host organism is called transgenic. Bacteria, plants, and animals have been genetically modified since the early 1970s for academic, medical, agricultural, and industrial purposes. Watch this short video explaining how scientists create a transgenic animal. Although the classic methods of studying the function of genes began with a given phenotype and determined the genetic basis of that phenotype, modern techniques allow researchers to start at the DNA sequence level and ask: “What does this gene or DNA element do?” This technique, called reverse genetics, has resulted in reversing the classical genetic methodology. One example of this method is analogous to damaging a body part to determine its function. An insect that loses a wing cannot fly, which means that the wing’s function is flight. The classic genetic method compares insects that cannot fly with insects that can fly, and observes that the non-flying insects have lost wings. Similarly in a reverse genetics approach, mutating or deleting genes provides researchers with clues about gene function. Alternately, reverse genetics can be used to cause a gene to overexpress itself to determine what phenotypic effects may occur. CRISPR Technology CRISPR stands for clustered regularly interspaced short palindromic repeats and represents a family of DNA sequences found within the genomes of prokaryotic organisms such as bacteria and archaea. These sequences are derived from DNA fragments of bacteriophages that have previously infected the prokaryote and are used to detect and destroy DNA from similar phages during subsequent infections. Hence these sequences play a key role in the antiviral defense system of prokaryotes. Figure \(9\) shows the crystal structure of a CRISPR RNA-guided surveillance complex, Cascade, bound to a ssDNA target, Cas9 (or "CRISPR-associated protein 9") is an enzyme that uses CRISPR sequences as a guide to recognize and cleave specific strands of DNA that are complementary to the CRISPR sequence. Cas9 enzymes together with CRISPR sequences form the basis of a technology known as CRISPR-Cas9 that can be used to edit genes within organisms. This editing process has a wide variety of applications including basic biological research, the development of biotechnology products, and the treatment of diseases. Figure \(10\)s shows a diagram of the CRISPR prokaryotic antiviral defense mechanism (10\): Diagram of the CRISPR prokaryotic antiviral defense mechanism. Image by James Atmos The CRISPR-Cas system is a prokaryotic immune system that confers resistance to foreign genetic elements such as those present within plasmids and phages that provide acquired immunity. RNA harboring the spacer sequence helps Cas (CRISPR-associated) proteins recognize and cut foreign pathogenic DNA. Other RNA-guided Cas proteins cut foreign RNA. CRISPR is found in approximately 50% of sequenced bacterial genomes and nearly 90% of sequenced archaea.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/09%3A_Investigating_DNA/9.02%3A_Bioinformatics.txt

Search Fundamentals of Biochemistry A DNA microarray (also commonly known as a DNA chip or biochip) is a collection of microscopic DNA spots attached to a solid surface. Scientists use DNA microarrays to measure the expression levels of large numbers of genes simultaneously or to genotype multiple regions of a genome. Each DNA spot contains picomoles (10−12 moles) of a specific DNA sequence, known as probes (or reporters or oligos). These can be a short section of a gene or other DNA element that is used to hybridize a cDNA or cRNA (also called anti-sense RNA) sample (called target) under high-stringency conditions. Probe-target hybridization is usually detected and quantified by the detection of fluorophore-, silver-, or chemiluminescence-labeled targets to determine the relative abundance of nucleic acid sequences in the target. The original nucleic acid arrays were macroarrays approximately 9 cm × 12 cm and the first computerized image-based analysis was published in 1981. It was invented by Patrick O. Brown. Figure \(1\) shows a schematic of DNA Microarrays. Within the organisms, genes are transcribed and spliced to produce mature mRNA transcripts (red). The mRNA is extracted from the organism and reverse transcriptase is used to copy the mRNA into stable ds-cDNA (blue). In microarrays, the ds-cDNA is fragmented and fluorescently labeled (orange). The labeled fragments bind to an ordered array of complementary oligonucleotides, and measurement of fluorescent intensity across the array indicates the abundance of a predetermined set of sequences. These sequences are typically specifically chosen to report on genes of interest within the organism's genome. The core principle behind microarrays is a hybridization between two DNA strands, the property of complementary nucleic acid sequences to specifically pair with each other by forming hydrogen bonds between complementary nucleotide base pairs. A high number of complementary base pairs in a nucleotide sequence means tighter non-covalent bonding between the two strands. After washing off non-specific bonding sequences, only strongly paired strands will remain hybridized. Fluorescently labeled target sequences that bind to a probe sequence generate a signal that depends on the hybridization conditions (such as temperature),and washing after hybridization. The total strength of the signal, from a spot (feature), depends upon the amount of target sample binding to the probes present on that spot. Microarrays use relative quantitation in which the intensity of a feature is compared to the intensity of the same feature under a different condition, and the identity of the feature is known by its position. Figure \(2\)s shows the hybridization of target and probe DNA on a microarray. Many types of arrays exist and the broadest distinction is whether they are spatially arranged on a surface or coded beads: • The traditional solid-phase array is a collection of orderly microscopic "spots", called features, each with thousands of identical and specific probes attached to a solid surface, such as glass, plastic, or silicon biochip (commonly known as a genome chip, DNA chip or gene array). Thousands of these features can be placed in known locations on a single DNA microarray. • The alternative bead array is a collection of microscopic polystyrene beads, each with a specific probe and a ratio of two or more dyes, which do not interfere with the fluorescent dyes used on the target sequence. DNA microarrays can be used to detect DNA (as in comparative genomic hybridization), or detect RNA (most commonly as cDNA after reverse transcription) that may or may not be translated into proteins. The process of measuring gene expression via cDNA is called expression analysis or expression profiling. Fabrication Microarrays can be manufactured in different ways, depending on the number of probes under examination, costs, customization requirements, and the type of scientific question being asked. Arrays from commercial vendors may have as few as 10 probes or as many as 5 million or more micrometer-scale probes. Spotted vs. in situ synthesized arrays Microarrays can be fabricated using a variety of technologies, including printing with fine-pointed pins onto glass slides, photolithography using pre-made masks, photolithography using dynamic micromirror devices, ink-jet printing, or electrochemistry on microelectrode arrays. In spotted microarrays, the probes are oligonucleotides, cDNA, or small fragments of PCR products that correspond to mRNAs. The probes are synthesized before deposition on the array surface and are then "spotted" onto glass. A common approach utilizes an array of fine pins or needles controlled by a robotic arm that is dipped into wells containing DNA probes and then depositing each probe at designated locations on the array surface. The resulting "grid" of probes represents the nucleic acid profiles of the prepared probes and is ready to receive complementary cDNA or cRNA "targets" derived from experimental or clinical samples. This technique is used by research scientists around the world to produce "in-house" printed microarrays from their labs. These arrays may be easily customized for each experiment, because researchers can choose the probes and printing locations on the arrays, synthesize the probes in their labs (or collaborating facility), and spot the arrays. They can then generate their own labeled samples for hybridization, hybridize the samples to the array, and finally scan the arrays with their equipment. This provides a relatively low-cost microarray that may be customized for each study and avoids the costs of purchasing often more expensive commercial arrays that may represent vast numbers of genes that are not of interest to the investigator. Publications exist which indicate in-house spotted microarrays may not provide the same level of sensitivity compared to commercial oligonucleotide arrays, possibly owing to the small batch sizes and reduced printing efficiencies when compared to industrial manufacturers of oligo arrays. In oligonucleotide microarrays, the probes are short sequences designed to match parts of the sequence of known or predicted open reading frames. Although oligonucleotide probes are often used in "spotted" microarrays, the term "oligonucleotide array" most often refers to a specific technique of manufacturing. Oligonucleotide arrays are produced by printing short oligonucleotide sequences designed to represent a single gene or family of gene splice-variants by synthesizing this sequence directly onto the array surface instead of depositing intact sequences. Sequences may be longer (60-mer probes such as the Agilent design) or shorter (25-mer probes produced by Affymetrix) depending on the desired purpose; longer probes are more specific to individual target genes, shorter probes may be spotted in higher density across the array and are cheaper to manufacture. One technique used to produce oligonucleotide arrays includes photolithographic synthesis (Affymetrix) on a silica substrate where light and light-sensitive masking agents are used to "build" a sequence one nucleotide at a time across the entire array. Each applicable probe is selectively "unmasked" before bathing the array in a solution of a single nucleotide, then a masking reaction takes place and the next set of probes are unmasked in preparation for a different nucleotide exposure. After many repetitions, the sequences of every probe become fully constructed. More recently, Maskless Array Synthesis from NimbleGen Systems has combined flexibility with large numbers of probes. Figure \(3\) shows a diagram of a typical dual-color microarray experiment. Two-color microarrays or two-channel microarrays are typically hybridized with cDNA prepared from two samples to be compared (e.g. diseased tissue versus healthy tissue) and that are labeled with two different fluorophores. Fluorescent dyes commonly used for cDNA labeling include Cy3, which has a fluorescence emission wavelength of 570 nm (corresponding to the green part of the light spectrum), and Cy5 with a fluorescence emission wavelength of 670 nm (corresponding to the red part of the light spectrum). The two Cy-labeled cDNA samples are mixed and hybridized to a single microarray that is then scanned in a microarray scanner to visualize the fluorescence of the two fluorophores after excitation with a laser beam of a defined wavelength. Relative intensities of each fluorophore may then be used in ratio-based analysis to identify up-regulated and down-regulated genes. Oligonucleotide microarrays often carry control probes designed to hybridize with RNA spike-ins. The degree of hybridization between the spike-ins and the control probes is used to normalize the hybridization measurements for the target probes. Although absolute levels of gene expression may be determined in the two-color array in rare instances, the relative differences in expression among different spots within a sample and between samples are the preferred method of data analysis for the two-color system. Examples of providers for such microarrays include Agilent with their Dual-Mode platform, Eppendorf with their DualChip platform for colorimetric Silverquant labeling, and TeleChem International with Arrayit. In single-channel microarrays or one-color microarrays, the arrays provide intensity data for each probe or probe set indicating a relative level of hybridization with the labeled target. However, they do not truly indicate abundance levels of a gene but rather a relative abundance when compared to other samples or conditions when processed in the same experiment. Each RNA molecule encounters protocol and batch-specific bias during the amplification, labeling, and hybridization phases of the experiment making comparisons between genes for the same microarray uninformative. The comparison of two conditions for the same gene requires two separate single-dye hybridizations. Several popular single-channel systems are the Affymetrix "Gene Chip", Illumina "Bead Chip", Agilent single-channel arrays, the Applied Microarrays "CodeLink" arrays, and the Eppendorf "DualChip & Silverquant". One strength of the single-dye system lies in the fact that an aberrant sample cannot affect the raw data derived from other samples, because each array chip is exposed to only one sample (as opposed to a two-color system in which a single low-quality sample may drastically impinge on overall data precision even if the other sample was of high quality). Another benefit is that data are more easily compared to arrays from different experiments as long as batch effects have been accounted for.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/09%3A_Investigating_DNA/9.04%3A_DNA_Microarrays.txt

Search Fundamentals of Biochemistry In situ hybridization (ISH) is a type of hybridization that uses a labeled complementary DNA, RNA, or modified nucleic acids strand (i.e., probe) to localize a specific DNA or RNA sequence in a portion or section of tissue (in situ) or if the tissue is small enough (e.g., plant seeds, Drosophila embryos), in the entire tissue (whole mount ISH), in cells, and in circulating tumor cells (CTCs). This is distinct from immunohistochemistry, which usually localizes proteins in tissue sections. In situ hybridization is used to reveal the location of specific nucleic acid sequences on chromosomes or in tissues, a crucial step for understanding the organization, regulation, and function of genes. The key techniques currently in use include in situ hybridization to mRNA with oligonucleotide and RNA probes (both radio-labeled and hapten-labeled), analysis with light and electron microscopes, whole mount in situ hybridization, double detection of RNAs and RNA plus protein, and fluorescent in situ hybridization to detect chromosomal sequences. DNA ISH can be used to determine the structure of chromosomes. Fluorescent DNA ISH (FISH) can, for example, be used in medical diagnostics to assess chromosomal integrity. RNA ISH (RNA in situ hybridization) is used to measure and localize RNAs (mRNAs, lncRNAs, and miRNAs) within tissue sections, cells, whole mounts, and circulating tumor cells (CTCs). In situ hybridization was invented by Mary-Lou Pardue and Joseph G. Gall. For hybridization histochemistry, sample cells and tissues are usually treated to fix the target transcripts in place and to increase access to the probe. As noted above, the probe is either a labeled complementary DNA or, now most commonly, a complementary RNA (riboprobe). The probe hybridizes to the target sequence at elevated temperature, and then the excess probe is washed away (after prior hydrolysis using RNase in the case of unhybridized, excess RNA probe). Solution parameters such as temperature, salt, and/or detergent concentration can be manipulated to remove any non-identical interactions (i.e., only exact sequence matches will remain bound). Then, the probe that was labeled with either radio-, fluorescent- or antigen-labeled bases (e.g., digoxigenin) is localized and quantified in the tissue, using either autoradiography, fluorescence microscopy, or immunohistochemistry, respectively. ISH can also use two or more probes, labeled with radioactivity or other non-radioactive labels, to simultaneously detect two or more transcripts. An alternative technology, branched DNA assay, can be used for RNA (mRNA, lncRNA, and miRNA) in situ hybridization assays with single molecule sensitivity without the use of radioactivity. This approach (e.g., ViewRNA assays) can be used to visualize up to four targets in one assay, and it uses patented probe design and bDNA signal amplification to generate sensitive and specific signals. Samples (cells, tissues, and CTCs) are fixed, then treated to allow RNA target accessibility (RNA un-masking). Target-specific probes hybridize to each target RNA. Subsequent signal amplification is predicated by specific hybridization of adjacent probes (individual oligonucleotides [oligos] that bind side by side on RNA targets). A typical target-specific probe will contain 40 oligonucleotides, resulting in 20 oligo pairs that bind side-by-side on the target for the detection of mRNA and lncRNA, and 2 oligos or a single pair for miRNA detection. Signal amplification is achieved via a series of sequential hybridization steps. A pre-amplifier molecule hybridizes to each oligo pair on the target-specific RNA, then multiple amplifier molecules hybridize to each pre-amplifier. Next, multiple-label probe oligonucleotides (conjugated to alkaline phosphatase or directly to fluorophores) hybridize to each amplifier molecule. A fully assembled signal amplification structure “Tree” has 400 binding sites for the label probes. When all target-specific probes bind to the target mRNA transcript, an 8,000-fold signal amplification occurs for that one transcript. Separate but compatible signal amplification systems enable multiplex assays. The signal can be visualized using a fluorescence or brightfield microscope.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/09%3A_Investigating_DNA/9.05%3A_In_Situ_Hybridization.txt

Search Fundamentals of Biochemistry Introduction to Lipids (Thanks to Rebecca Roston for providing a cohesive organizational framework and image templates) Lipids are organic molecule molecules that are soluble in organic solvents, such as chloroform/methanol, but sparingly soluble in aqueous solutions. These solubility properties arise since lipids are mostly hydrophobic. One type, triglycerides, is used for energy storage since they are highly reduced and get oxidized to release energy. Their hydrophobic nature allows them to pack efficiently through self-association in an aqueous environment. Triglycerides are also used for insulation since they conduct heat poorly, which is good if you live in a cold climate but bad if you wish to dissipate heat in a hot one. Triglycerides also offer padding and mechanical protection from shocks (think walruses). Another type of lipid forms membrane bilayers, which separate cellular contents from the outside environment, or separate intracellular compartments (organelles) from the cytoplasm. Some lipids are released from cells to signal other cells to change to specific stimuli in a process called cell signaling. From a more molecular perspective, lipids can act as cofactors for enzymes, pigments, antioxidants, and water repellents. As we saw with proteins, lipid structure mediates their function. So let's probe their structures. Lipids can be split into structural classes in a variety of ways. An earlier classification divided them into those that release fatty acids on based-catalyzed hydrolysis to form soaps in a saponification reaction and those that don't. A much better and broader classification is based on if the lipids contain fatty acids or isoprenoids, as shown in Figure \(1\) below. Figure \(1\): Fatty acids and isoprenoid lipids The nonpolar chains of the fatty acid are drawn in the figure above in the lowest energy zig-zag fashion as we saw when we discussed the main chain conformation of proteins (Chapter 4.1). In that chapter, we started with the exploration of a long 12 C chain carboxylic acid, dodecanoic acid. In the lowest energy conformation, the dihedral angles are all + 1800 to minimize torsional strain in the molecule. Rotation around one C-C bond can produce a gauche form, which introduces a kink into the chain as shown in Figure \(2\). Fatty Acids Fatty acids can be free or covalently linked by ester or amide link to a base molecule like triglycerides or membrane lipids. The key principle that we learned with our study of proteins, that structure determines function, also applies to lipids. The figure below shows three different types of molecules, a free fatty acid, a wax with an esterified fatty acid, and a glycolipid with a fatty acid connected by an amide link in another type of lipid (glycosphingolipid). Each has different properties leading to different functions. Waxes for instance are very nonpolar and water-insoluble. They are amorphous solids at room temperature but, depending on their structure, can easily melt to form high-viscosity liquids. They are used as a coating on the surfaces of leaves to help prevent water loss. The glycolipids (glyco- as it contains a monosaccharide group) are constituents of membrane bilayers. Figure \(3\) shows the general structures of fatty acid-containing waxes and other lipids. Fatty acids vary in length, usually contain an even number of carbon atoms, and can be saturated (have no double bonds in the acyl chain), or unsaturated (with either one -monounsaturated - or multiple - polyunsaturated - cis double bond(s)). The double bonds are NOT conjugated as they are separated by a methylene (-CH2-) spacer. Fatty acids can be named in many ways. • symbolic name: given as x:y Δ a,b,c where x is the number of carbon atoms in the chain, y is the number of double bonds, and a, b, and c are the positions of the start of the double bonds counting from C1 - the carboxyl carbon. Double bonds are usually cis (Z). • systematic name using IUPAC nomenclature. The systematic name gives the number of carbon atoms in the chain (e.g. hexadecanoic acid for 16:0). If the fatty acid is unsaturated, the base name reflects the number of double bonds (e.g. octadecenoic acid for 18:1 Δ 9 and octadecatrienoic acid for 18:3Δ 9,12,15). • common name: (e.g. oleic acid, 18:1Δ9), which is found in high concentration in olive oil) They can be named most easily with a symbolic name. Figure \(4\) shows examples of fatty acids and their symbolic names. There is an alternative to the symbolic representation of fatty acids, in which the carbon atoms are numbered from the distal end (the n or ω end) of the acyl chain (the opposite end from the alpha carbon). Hence 18:3 Δ 9,12,15 could be written as 18:3 (ω -3) or 18:3 (n -3), where the terminal C is numbered one and the first double bond starts at C3. The most common saturated fatty acids found in biochemistry textbooks are listed in the table below. Note how the melting point increases with the length of the hydrocarbon chain. This arises from increasing noncovalent induced dipole-induced dipole attractions between the long chains. Heat must be added to lessen these attractions to allow melting. Table \(1\) below shows examples of fatty acids and their symbolic names. Symbolic common name systematic name structure mp(C) 12:0 Lauric acid dodecanoic acid CH3(CH2)10COOH 44.2 14:0 Myristic acid tetradecanoic acid CH3(CH2)12COOH 52 16:0 Palmitic acid Hexadecanoic acid CH3(CH2)14COOH 63.1 18:0 Stearic acid Octadecanoic acid CH3(CH2)16COOH 69.6 20:0 Arachidic aicd Eicosanoic acid CH3(CH2)18COOH 75.4 Table \(1\): Examples of fatty acids and their symbolic names Table \(2\) below shows common unsaturated fatty acids. Arachidonic acid is an (ω -6) fatty acid while docosahexaenoic acid is an (ω -3) fatty acid. Note the decreasing melting point for the 18:X series with an increasing number of double bonds. Symbol common name systematic name structure mp(C) 16:1Δ9 Palmitoleic acid Hexadecenoic acid CH3(CH2)5CH=CH-(CH2)7COOH -0.5 18:1Δ9 Oleic acid 9-Octadecenoic acid CH3(CH2)7CH=CH-(CH2)7COOH 13.4 18:2Δ9,12 Linoleic acid 9,12 -Octadecadienoic acid CH3(CH2)4(CH=CHCH2)2(CH2)6COOH -9 18:3Δ9,12,15 α-Linolenic acid 9,12,15 -Octadecatrienoic acid CH3CH2(CH=CHCH2)3(CH2)6COOH -17 20:4Δ5,8,11,14 arachidonic acid 5,8,11,14- Eicosatetraenoic acid CH3(CH2)4(CH=CHCH2)4(CH2)2COOH -49 20:5Δ5,8,11,14,17 EPA 5,8,11,14,17-Eicosapentaenoic- acid CH3CH2(CH=CHCH2)5(CH2)2COOH -54 22:6Δ4,7,10,13,16,19 DHA Docosohexaenoic acid 22:6w3 Table \(2\): Common unsaturated fatty acids Let's consider how the presence of double bonds in fatty acids influences their melting points. Figure \(5\) shows common variants of fatty acids each with 18 carbon atoms. Compare the Lewis structure and spacefill models below. What a difference a cis double bond makes! The double bonds in fatty acids are cis (Z), which introduces a "permanent" kink into the chain, similar to the"temporary" kink in the saturated dodecanoic acid carboxylic acid with a single gauche bond (Figure 2). The kink is permanent since there is no rotation around the double bond unless it is broken (which can happen through photoisomerization reactions). The more double bonds, the greater the kinking. The more kinks, the less chance for van der Waals contacts between the acyl chains, and reduces induced-dipole-induced dipole interactions between the chains, leading to lowered melting points. Now in your mind, replace the cis double bond in oleic acid with a trans. You should now see a long "zig-zag" shaped molecule with no kinks. Trans fatty acids are rare in biology but are produced in the industrial partial hydrogenation of fats, which is done to decrease the number of double bonds and make the fats more solid-like and tastier while decreasing rancidity. These trans fatty acids would pack closer together and affect the structure and function of the lipid in a given environment (such as a membrane bilayer). Increased consumption of trans fatty acids is associated with an increased risk of cardiovascular disease. Table \(3\): shows the percentages of fatty acids in different oils/fats FAT <16:0 16:1 18:0 18:1 18:2 18:3 20:0 22:1 22:2 . Coconut 87 . 3 7 2 . . . . . Canola 3 .   11 13 10 . 7 50 2 Olive Oil 11 . 4 71 11 1 . . . . Butter-fat 50 4 12 26 4 1 2 . . . Table \(3\): Percentages of fatty acids in different oils/fats The fatty acid composition differs in different organisms: • animals have 5-7% of fatty acids with 20-22 carbons, while fish have 25-30% • animals have <1% of their fatty acids with 5-6 double bonds, while plants have 5-6% and fish 15-30% As there are essential amino acids that cannot be synthesized by humans, there are also essential fatty acids that must be supplied by the diet. There are only two, one each in the n-6 and n-3 classes: • n-6 class: α-linoleic acid (18:2 n-6, or 18:2Δ9,12) is a biosynthetic precursor of arachidonic acid (20:4 n-6 or 20:4Δ5,8,11,14) • n-3 class: linolenic acid (18:3 n-3, or 18:3Δ9,12,15) is a biosynthetic precursor of eicosapentaenoic acid (EPA, 20:5 n-3 or 20:5Δ5,8,11,14,17) and to a much smaller extent, docosahexaenoic acid (DHA, 22:6 n-3 or 22:6Δ4,7,10,13,16,19). These two fatty acids are essential since mammals cannot introduce double bonds in fatty acids beyond carbon 9. These essential precursor fatty acids are substrates for intracellular enzymes such as elongases and desaturases (to produce 20:4 n-6, 20:5 n-3, and 22:6 n-3 fatty acids), and beta-oxidation type enzymes in the endoplasmic reticulum and another organelle, the peroxisome. The peroxisome is involved in the oxidative metabolism of straight-chain and branched fatty acids, peroxide metabolism, and cholesterol/bile salt synthesis. Animals fed diets high in plant 18:2(n-6) fats accumulate 20:4(n-6) fatty acids in their tissues while those fed diets high in plant 18:3(n-3) accumulate 22:6(n-3). Animals fed diets high in fish oils accumulate 20:5 (EPA) and 22:6 (DHA) at the expense of 20:4(n-6). Many studies support the claim that diets high in fish that contain abundant ω-3 fatty acids, in particular EPA and DHA, reduce inflammation and cardiovascular disease. ω-3 fatty acids are abundant in high-oil fish (salmon, tuna, sardines), and lower in cod, flounder, snapper, shark, and tilapia. Some suggest that contrary to images of early hominids as hunters and scavengers of meat, human brain development might have required the consumption of fish which is highly enriched in arachidonic and docosahexaenoic acids. A large percentage of the brain consists of lipids, which are highly enriched in these two fatty acids. These fatty acids are necessary for the proper development of the human brain and in adults. Deficiencies in these might contribute to ADHD, dementia, and dyslexia. These fatty acids are essential in the diet and probably could not have been derived in high enough amounts from the eating of the brains of other animals. The mechanism for the protective effects of n-3 fatty acids in health will be explored later in the course when we discuss prostaglandins synthesis and signal transduction. Aggregates of Fatty Acids in Aqueous Solution: Micelles Structure determines both properties and function. It should be obvious that free, unesterified fatty acids are very (but not completely) insoluble in water. When added to water, they saturate the solution at a very low concentration and then phase separate out into aggregates called micelles. The structure of a micelle formed from dodecylsulfate, a common detergent with a sulfate instead of a carboxylate head group, is shown below. All of the nonpolar Cs and Hs of the long alkyl chains are "buried" and are not exposed to water, whereas the sulfate head groups are solvent exposed. Figure \(6\) shows an interactive iCn3D model of image Figure \(6\): Sodium dodecylsulfate micelle (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...KECTiGaPPScED7 Fatty acids (carboxylates and sulfates) are amphiphilic, with a larger polar/charged head group that tapers down to a hydrophobic tail, forming a cone-like structure. This cone structure allows the packing of many of these single chains amphiphiles into a micelle, as shown in Figure \(7\). ​Free fatty acids are transported in the cell and the blood not in micelles but by fatty acid-binding proteins. The most abundant protein in the blood, albumin, binds and transports fatty acids. Figure \(8\) shows an interactive iCn3D model of hexadecanoic acid bound to human albumin (1E7H). The fatty acids are shown in colored spacefill rendering. Figure \(8\): Hexadecanoic acid bound to human albumin (1E7H). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...CGUdMj11vNevF6 Waxes You are familiar with ear wax and also the waxy surface of plants. Ear wax contains long-chain fatty acids (saturated and unsaturated) and alcohols derived from them. (They also contain isoprenoid derivatives like squalene and cholesterol). The waxy cuticle surface layers of plants contain very-long-chain fatty acids ( C20–C34) and their derivatives including alkanes, aldehydes, primary and secondary alcohols, ketones, and esters, (They also contain isoprenoid derivatives as well). We will consider waxes as very long-chain fatty acids and their derivatives, as shown in Figure \(9\). These molecules are extremely nonpolar and as such makes great barriers preventing water loss through leaves and water penetration into the ear. This group of molecules clearly shows that the properties (insolubility, high melting point) and function (hydrophobic barrier/protection) arises from structure (very long chain carbon molecules, few electronegative atoms, and lack of C=C double bonds). Fatty Acid-Containing Lipids We can categorize these lipids based on function or structure (even though these are related). Function • storage lipids - triacylglycerols • membrane lipids - many different lipids Structure • glycerolipids, which use glycerol as a backbone for fatty acid attachment • sphingolipids, which use sphingosine as a backbone The structures of glycerol and sphingosine are shown in Figure \(10\). Fatty acids are connected to these two "backbone" structures by either ester (mostly) or amide links. Let's explore the classes of fatty acid-containing lipids that use these two "backbone" structures. Storage Lipids - Triacylglycerols (TAGs) Triacylglycerols contain the majority of fatty acids in species that store fatty acids for energy. You will often see them named triglycerides or triacylglyceride, but this is a term used more in clinical chemistry and industry (and often in the media). Figure \(11\) shows a schematic diagram of the glycerol backbone with three fatty acids esterified to it. They are glycerolipids as they contain a glycerol base. Glycerol is not chiral but given the incredible diversity of fatty acids, glycerols likely have three different fatty acids esterified to them, making them chiral. If triacylglycerols contain predominately saturated fatty acids, they are solids at room temperature and are called fats. Those with multiple double bonds in the fatty acids are likely liquids at room temperature. These are called oils. Triacylglycerols are even more insoluble than fatty acids, which contain a polar and mostly charged carboxylate. Figure \(12\) shows a triacylglycerol containing all 16:0 saturated fatty acids (left) and one containing all 16:2Δ9,12 polyunsaturated fatty acids. The figures were constructed with a specific set of dihedral angles to illustrate a point, that polyunsaturated fats in a triacylglycerol don't pack as tightly and have lower induced dipole-induced dipole attractions between the acyl chains than is possible with saturated fatty acyl chains. Hence the melting point of triacylglycerols containing polyunsaturated fatty acids is lower than for those with saturated ones. Let's repeat the key mantra: the structure of lipids determines their function. Consider the very insoluble triacylglycerols which are used as the predominant storage form of chemical energy in the body. In contrast to polysaccharides such as glycogen (a polymer of glucose), the carbon atoms in the acyl chains of the triacylglycerol are in a highly reduced state. The main source of energy to drive not only our bodies but also our society is obtained through oxidizing carbon-based molecules to carbon dioxide and water, in a reaction that is highly exergonic and exothermic. Sugars are already part way down the free energy "hill" since each carbon is partially oxidized. 9 kcal/mol (38 kJ/mol) can be derived from the complete oxidation of fats, in contrast to 4.5 kcal/mol (19 kJ/mol) from that of proteins or carbohydrates. In addition, glycogen is highly hydrated. For every 1 g of glycogen, 2 grams of water is H-bonded to it. Hence it would take 3 times more weight to store the equivalent mass of carbohydrates compared to triacylglycerol, which is stored in anhydrous lipid "droplets" within cells. In addition, fats are more flexible, given the large number of conformations available to the acyl chain C-C bonds by simple rotation around the C-C bonds. Polysaccharides have monomeric cyclohexane-like chair structures and are much more rigid. Another interesting point is that glucose and glycogen are found in cells, and they can be mobilized quickly for energy needs. Yes, fats are present in all cells as well (for example all cells have interior and exterior cell membranes). However, the major storage form of fat, triacylglycerols, is stored in special cells called adipocytes, which comprise adipose or fat tissue, and must be mobilized by signaling agents and transported in the form of fatty acids to cells for utilization. Again, triacylglycerols don't form membranes, which separate outside and inside aqueous environments. They are simply so insoluble that they phase-separate into lipid droplets. Their formation and structure are a bit more complex than that, though, and we will discuss lipid droplets more in the next section. Membrane Lipids Membranes are bilayers of amphiphilic lipids that separate the outside and inside (cytoplasm) aqueous environments in cells and the cytoplasm and interior contents of organelles within cells. In general, single-chain lipid amphiphiles form micelles. Amphiphilic membrane lipids typically have two nonpolar tails connected to a polar head, giving them a less conical and more cylindrical shape that disallows micelle formation while favoring bilayer formation. Figure \(13\) shows a short section of a bilayer membrane made from lipids with a polar (and charged) head group (phosphocholine) and two 16:0 chains. The red and blue spheres represent the O and N atoms of the head groups, which are sequestered to the exterior parts of the bilayer, where they would interact with water. The nonpolar 16:0 tails are shown in cyan, clearly illustrating the nonpolar nature of the interior of the bilayer. Figure \(14\) shows an interactive iCn3D model of a hydrated bilayer of the di16:0 phosphatidycholine bilayer. Figure \(14\): Hydrated di16:0 phosphatidycholine bilayer (Copyright; author via source). Click the image for a popup. We will consider two general types of fatty acid-containing membrane lipids, glycerolipids, with two fatty acids esterified to a glycerol base, and sphingolipids with one fatty acid in amide link to a different base, sphingosine. Sphingosine comes with its own built-in long alkyl chain that provides the "second" nonpolar chain. There are many different polar/charged head groups for these membrane lipids. Now let's look in more detail at the double-chain amphiphiles comprising these membrane bilayers. Glycerolipids There are two main types of glycerolipids, glycerophospholipids, and glyceroglycolipids, which are the most common lipids in membranes. Glycerophospholipids Figure \(15\) shows the structural features and nomenclature for glycerophospholipids. These lipids have enormous structural variability given the large number of different fatty acids (both saturated and unsaturated) and head groups that can be attached to a phosphate attached to the carbon 3 of glycerol. The structures of the most common glycerophospholipids are shown in Figure \(16\). Phosphatidylcholine (PC) has the common name lecithin while phosphatidylserine (PS) is called cephalin. Note that the head groups all have charges since they all have a negatively charged phosphate. PS has two additional charged atoms which would effectively cancel out. PE has a charged amine but could become uncharged at pH values approaching its pKa. PC has a quaternary amine which is charged independent of pH, which would give PC a net 0 charge but with two discrete charges. Glyceroglycolipids These do not have a phosphate group attached to the oxygen on C3 of glycerol. Rather they have a mono- or oligosaccharide or, more loosely, a betaine group, each attached by an ether linkage to the glycerol C3 carbon. Figure \(17\) Structural features and nomenclature for glyceroglycolipids. Figure \(18\) shows some examples of glyceroglycolipids. Again there are an enormous number of different glycoglycerolipids, owing to the diversity of head groups and fatty acids esterified to glycerol at C1 and C2. The above figure (right) shows an example that has no phosphate but also doesn't have a mono- or polysaccharide for a head group. Rather it has a betaine group. Betaine is the common name for trimethylglycine but is used for any N-trimethylated amino acids. Betaine glycerolipids are found in lower eukaryotic organisms (algae, fungi, and some protozoa), in photosynthetic bacteria, and in some spore-producing plants like ferns. Some would call these lipoamino acids. Membrane Lipids - Sphingolipids Sphingolipids contain a sphingosine backbone and a fatty acid linked through an amide bound. The simplest sphingolipids are the ceramides, which are double-chain amphiphiles but without further modification of the functional groups on the polar head of sphingosine. The structures of both sphingosine and ceramide are shown below in Figure \(19\): Figure \(19\): Structures of sphingosine and ceramide Ceramides are abundant in the skin where they help provide a protective barrier. Skin creams also contain ceramides. These lipids make up only about 1% of body lipids and there are over 200 different ceramides in humans arising from the different fatty acids linked to sphingosine. They have a key role in cell signaling and can also affect cardiovascular health in ways that are just being appreciated. Since they are double-chain amphiphiles, they are found in membranes where they alter the properties of the bilayer (including membrane fluidity). Ceramides with long acyl tails (especially those with 16, 18, or 24 carbons) seem especially deleterious. Phosphosphingolipids and glycosphingolipids These membrane lipids have a ceramide base but also contain modifications at the polar functional groups of the sphingosine head. Let's consider the phosphosphingolipids and the glycosphingolipids together. These groups do not use glycerol as a base for the attachment of fatty acids and head groups. Rather they use molecule sphingosine. Figure \(20\) shows the structural features and nomenclature of sphingolipids. Examples of both classes are shown in Figure \(21\). Note the base sphingosine (in red) provides an amine to attach a fatty acid through an amide bond and an OH for attachment of the head group. Sugar-containing glycosphingolipids are found largely in the outer face of plasma membranes. The primary lipid of myelin, which coats neuronal axons and insulates them from loss of electrical signaling down the axon, is galactocerebroside Figure \(22\) shows a summary of all of the different types of fatty acid-containing lipids Over a 1000 different lipids are found in eukaryotic cells. This complexity has led to the development of an even more comprehensive classification system for lipids. In this system, lipids are given a very detailed as well as all-encompassing definition: "hydrophobic or amphipathic small molecules that may originate entirely or in part by carbanion-based condensations of thioesters (fatty acyl, glycerolipids, glycerophospholipids, sphingolipids, saccharolipids, and polyketides) and/or by carbocation-based condensations of isoprene units (prenol lipids and sterol lipids)." Eight different categories of lipids are listed in the parentheses above. We will stick to the definition used throughout this chapter. Shapes of membrane lipids Let's look at the general shape of the double-chain amphiphiles that make bilayers. We saw the long-chain fatty or sulfate acids form conical structures which fit nicely together when they self-aggregate to form micelles. In contrast, membrane-forming double-chain amphiphiles have more cylindrical shapes that can't be fitted together in micelles but rather form a less curved bilayer structure, as shown in Figure \(23\). We will consider the variety of membrane structures in the next section. Triacylglyceride/Phospholipid Stereochemistry Glycerol is an achiral molecule since C2 has two identical substituents, -CH2OH. Glycerol in the body can be chemically converted to triglycerols and phospholipids (PL) which are chiral, and exist in one enantiomeric form. How can this be possible if the two CH2OH groups on C2 of glycerol are identical? It turns out that even though these groups are stereochemically equivalent, we can differentiate them as described in the figure below. Let's replace the -CH2OH in one of the end carbons with -CH2OD. With this simple change, glycerol is now chiral. Look at the top half of Figure \(24\). Glycerol is oriented with the OH on C2 (the middle carbon) pointing to the left. The OH of the top carbon in this orientation, C1, is replaced with OD, where D is deuterium to make the molecule chiral (four different groups attached to C2). By rotating the molecule such that the H on C2 points to the back, and assigning priorities to the other substituents on C2 (OH =1, DOCH2 =2, and CH2OH = 3), it can be seen that the resulting molecule is in the S configuration. We simply name the C1 carbon which we modified with deuterium as the proS carbon. Likewise, if we replaced the OH on C3 with OD, we will form the R enantiomer. Hence C3 is the proR carbon. This shows that in reality, we can differentiate between the two identical CH2OH substituents. We say that glycerol is not chiral, but prochiral. (Think of this as glycerol has the potential to become chiral by modifying one of two identical substituents.) In the bottom half of Figure \(23\), we can relate the configuration of glycerol above, (when OH on C2 is pointing to the left) to the absolute configuration of L-glyceraldehyde, a simple sugar (a polyhydroxyaldehyde or ketone), another 3C glycerol derivative. This molecule is chiral with the OH on C2 (the only chiral carbon) pointing to the left. It is easy to remember that any L sugar has the OH on the Last chiral carbon pointing to the Left. The enantiomer (mirror image isomer) of L-glyceraldehyde is D-glyceraldehyde, in which the OH on C2 points to the right. Biochemists use L and D for lipid, sugar, and amino acid stereochemistry, instead of the R, S nomenclature you used in organic chemistry. The stereochemical designation of all the sugars, amino acids, and glycerolipids can be determined from the absolute configuration of L- and D-glyceraldehyde. Now let's see how an enzyme can take a prochiral molecule like glycerol and phosphorylate only one of the -CH2OHs to make one specific isomer, glycerol-3-phosphate, a key intermediate in the biosynthesis of phosphatidic acid (PA), a glycerophospholipid, as well as chiral triacylglycerols, shown in Figure \(25\). The far left part of the pathway shows how the proR CH2OH of glycerol is phosphorylated to produce one specific enantiomer, L-glycerol-3-phosphate. (The top part of the figure shows another way to make this molecule from glucose through the glycolytic pathway we will encounter in a future chapter. The first step (above figure) involves the phosphorylation of the OH on C3 by ATP (a phosphoanhydride similar in structure to acetic anhydride, an excellent acetylating agent) to produce the chiral molecule glycerol phosphate. Based on the absolute configuration of L-glyceraldehyde, and using this to draw glycerol (with the OH on C2 pointing to the left), we can see that the phosphorylated molecule can be named L-glycerol-3-phosphate. However, by rotating this molecule 180 degrees, without changing the stereochemistry of the molecule, we don't change the molecule at all, but using the D/L nomenclature above, we would name the rotated molecule D-glycerol-1-phosphate. This is illustrated in Figure \(26\). We can’t give the same molecule two different names. Hence biochemists have developed the stereospecific numbering system (sn), which assigns the 1-position of a prochiral molecule to the group occupying the proS position. The proS C1 is hence at the sn-1 position. With that designation, C2 is at the sn-2 position, and C3 is at the sn-3 position. Using this nomenclature, we can see that the chiral molecule described above, glycerol-phosphate, can be unambiguously named as sn-glycerol-3-phosphate. The hydroxyl substituent on the proR carbon was phosphorylated. It is interesting to note that archaea use isoprenoid chains linked by ether bonds to sn-glycerol 1-phosphate in their synthetic pathways. As noted above, bacteria and eukaryotes use fatty acids attached by ester bonds to sn-glycerol 3-phosphate The enzymatic phosphorylation of the proR CH2OH of glycerol to form sn-glycerol-3-phosphate is illustrated in Figure \(27\). As we were able to differentiate the 2 identical CH2OH substituents as containing either the proS or proR carbons, so can the enzyme. The enzyme can differentiate identical substituents on a prochiral molecule if the prochiral molecule interacts with the enzyme at three points. Another example of a prochiral reactants/enzyme system involves the oxidation of the prochiral molecule ethanol by the enzyme alcohol dehydrogenase, in which only the proR H of the 2 H’s on C2 is removed. (We will discuss this later.) Isoprenoid-containing lipids This is the last class of lipids we will consider. They do not contain fatty acids. Rather they contain isoprene, a small branched alkadiene, which can polymerize into larger molecules containing isoprene monomer to form isoprenoids, often called terpenes. Instead of using isoprene as the polymerization monomer, either dimethylallyl pyrophosphate (DMAPP) or isopentenylpyrophosphate (IPP) are used biologically. Figure \(28\) shows how DMAPP and IPP (both containing 5Cs) are used in a polymerization reaction to form geranyl-pyrophosphate (C10), farnesyl pyrophosphate (C15) and geranyl-geranyl pyrophosphate (C20). Many isoprenoid lipids are made from farnesyl pyrophosphate. For membrane purposes, the most important of these is cholesterol. Figure \(29\) shows an overview of the synthesis of cholesterol from two farnesyl pyrophosphates linking together in a "tail-to-tail" reaction to form squalene, a precursor of cholesterol. Each isoprene unit (5Cs) is shown in different colors to make it easier to see. Other biologically important isoprenoid-containing vitamins are shown in Figure \(30\). The smell of fresh rain Everyone who has walked in the woods after a fresh rain knows the smell of a terpene called geosmin, "an Earthy-smelling substance" found in abundance in a group of substances collectively called petrichor, derived from the Greek petra (rock) and ichor (blood of gods). The structure of geosmin is shown below. These substances are released when water falls or soils and rocks, and seeps into pores from which aerosols are released. Too hard of a rain will saturate pores in rocks and prevent the release. Some insects like the springtail are attracted to geosmin. The synthesis of geosmin by bacteria and blue-green algae is under the control of transcription factors which also affect bacterial spore formation. The attracted insects carry away the spores. Flies are repelled by geosmin, which is detected by a receptor at very low geosmin concentration. In contrast, the Aedes aegypti mosquito is attracted to geosmin which parallels the fact that the mosquito is not affected by bacterial toxins. One last look at lipid structure and shapes With the exclusion of waxes and triacylglycerols, the other lipids we have discussed, including the mostly planar molecule cholesterol, are amphiphilic. We have seen that single-chain fatty acids form micelles while lipids with two nonpolar chains and a polar/charged head group form bilayers. Given the relative size of the head group and the degree of unsaturation of the double bonds in fatty acids, the overall shapes of the membrane-forming lipids differ as illustrated below. They are arranged like dominos in the membrane based on their geometric volumes. Preferential clustering of identical types can cause local and extended changes in a prototypical bilayer structure. Membranes and their components must be dynamic to enable all the functions and activities of a membrane. Ligands bind to membrane receptors (usually proteins), which can invaginate and pinch off to form an intracellular vesicle containing the receptor for processing. Likewise, vesicles can pinch off into the extracellular space. Cells must divide. Think of the membrane changes necessary for that! Also, consider that the length of cholesterol is half that of a typical double-chain amphiphile so it fits into just one of the bilayer lipid leaflets where it modulates lipid bilayer property. Figure \(31\) shows a series of lipids and their shape profiles. We will consider membranes in greater detail in section 10.3. Next, however, we will explore in more detail the properties of micelle and lipid droplet systems before addressing the more structurally complicated lipid bilayers. End-of-Chapter Questions Exercise \(1\) x) A cell membrane has the ability to remodel in response to stress to promote membrane integrity. In the situations below, how could the membrane be remodeled to prevent damage? (I.e. what types of lipids could be added/removed to ensure homeostasis) a) Increase in temperature b) x) What are the three classes of lipids? Explain their similarities and differences. x) For each lipid below, name the type of lipid (membrane lipid, triacylglycerol, storage lipid, sphingolipid, wax, sterol, membrane glycerolipid, none of these), if it could be found in membrane, and if it is fatty acid or isoprene derived. (Insert pics of lipids) Answer Add texts here. Do not delete this text first.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/10%3A_Lipids/10.01%3A_Introduction_to_lipids.txt

Search Fundamentals of Biochemistry Single Chain Amphiphiles and Micelles An understanding of lipids in simple solutions in the lab is incredibly helpful to understanding them in vivo. The same physical-chemical constraints would apply to the complex environment of the cell. What is different in the cell is that lipids are found in a cellular environment that is incredibly crowded with proteins that bind, synthesize, and breakdown lipids. Nevertheless, we can apply what we know from the test tube experiments to the cell. To understand how molecules might react, it helps a bit to pretend you are a molecule and ask yourself what would you do! We want to know how lipid molecules, specifically single and double-chain amphiphiles, interact with each other and solvent when they are added to water. Before you read the answer, look at the image below and ask yourself the question: What would I do if I were a single chain amphiphile and jumped into water as shown in Figure $1$? Here is what they do. When added to water, some single-chain amphiphiles dissolve in water while others form a monolayer on the surface of the water. If enough enter the solution and exceed their net solubility, they self-aggregate to form micelles. These outcomes are shown in Figure $2$. Figure $3$ shows an interactive iCn3D model of an sodium dodecylsulfate (SDS) micelle Figure $3$: Sodium dodecylsulfate (SDS) micelle (Copyright; author via source). Click the image for a popup or use this external link: not available Double-chain amphiphiles, in contrast, form bilayers instead of micelles. (Note: single and double-chain amphiphiles can form other multimolecular aggregate structures as well, but micelles and bilayers are the most common and are the only ones we will consider.) The micelle interior is completely nonpolar. Spherical bilayers that enclose an aqueous compartment are called vesicles or liposomes. Micelles and bilayers, formed from single and double-chain amphiphiles, respectively, represent noncovalent aggregates and hence are formed by an entirely physical process. No covalent steps are required. Common single-chain amphiphiles that form micelles are detergents (like sodium dodecyl sulfate - SDS) as well as fatty acids, which themselves are detergents. Sodium hydroxide feels slippery on your skin since the base hydrolyses the fatty acids esterified to skin lipids. The free fatty acids then aggregate spontaneously to form micelles which act like detergents, and are also slippery. Micelle/detergents in water are an example of an emulsion of two liquids that are generally immiscible in each other unless one is dispersed into small droplets into the other Fine oil drops can be dispersed in water and fine aqueous drops can be dispersed in a nonpolar liquid. Many vaccines are formulated as this later type of emulsion. Grease and oil in your clothes can be carried away by "diving" into the nonpolar part of the detergent micelle which is dispersed in water as an emulsion. Another example of an emulsion or more properly a colloid is a cloud, a dispersion of liquid water droplets in a solvent, the atmosphere. The formation of these structures can be understood from the study of noncovalent interactions but also through thermodynamics. In a micelle, the buried acyl chains can interact and be stabilized by induced dipole-induced dipole forces as the nonpolar carbons and hydrogen are in van der Walls contact. They are sequestered from water. This view fits our simple axiom of "like-dissolves like". The polar head groups can be stabilized by ion-dipole bonds between charged head groups and water. Likewise, H-bonds between water and the head group stabilize the exposed head groups in water. Repulsive forces may also be involved. Head groups can repel each other through steric factors, or ion-ion repulsion from like-charged head groups. The attractive forces must be greater than the repulsive forces, which lead to these molecular aggregates. From a thermodynamic approach, one problem arises with this simple explanation. For a micelle or bilayer to form, many monomers must aggregate to form a single micelle or vesicle, which is entropically disfavored! So let's delve into the thermodynamics of micelle formation. ΔG, the free energy chance for a reaction, determines the spontaneity and extent of a chemical or physical reaction. The free energy of the system depends on 3 variables, temperature T, pressure P, and n, the number of moles of each substance. For the latter, think of solute X on two different sides of a permeable membrane. If the concentration of X is the same on each side, as shown in the system below, the system is in equilibrium as shown in Figure $4$. If the system is composed of two different parts, A and B, the system is at equilibrium (ΔG=0) if TA = TB, PA = PB, and the change in the absolute free energy per mole of A is ΔGA/Δn = ΔGB/Δn. More precisely, using simple calculus, we would discuss incremental changes in absolute free energy/mol, dGA/dn for A (often called the chemical potential of A, μA), and dGB/dn or μB)for B. At equilibrium dGA/dn = dGB/dn. (We will use the symbol G here instead of μ). G then is the absolute free energy/mol (again often called the chemical potential), where G=Go +RTln[A]. The equations you used in introductory chemistry can be written. \begin{array}{l} \Delta \mathrm{G}=\Delta \mathrm{G}^{0}+\mathrm{RTIn} \mathrm{Qr} \ \Delta \mathrm{G}=\Delta \mathrm{H}-\mathrm{T} \Delta \mathrm{S} \ \Delta \mathrm{G}^{0}=\Delta \mathrm{H}^{0}-\mathrm{T} \Delta \mathrm{S}^{0} \ \Delta \mathrm{G}^{0}=-\mathrm{RTInK} \mathrm{eq} \end{array} Now let's apply this to the chemical equation for micelle formation: n SCA <==> 1 micelle where SCA represents a single chain amphiphile. At first glance, we might suspect that: • ΔH0 < 0 since the induced dipole-induced dipole interactions among the buried acyl chains in the micelle would be much more favorable than the water-acyl interactions for the monomeric amphiphile in solution. This notion is supported by our aphorism, "like dissolves like". Of course, we couldn't ignore polar interactions (H bonding for example) among the head groups and water, but we might expect these to be equally favorable in both the monomeric and micellular states. • ΔS0 < 0 since we are forming a very ordered state (a single micelle) with much less entropy from a state (single chains amphiphiles dispersed in solution) with much more entropy. Hence it would appear that micelle formation is enthalpically favored but entropically disfavored. Let's examine this issue more closely. First, we need to obtain a greater understanding of ΔGo which should give us a clue as to where a SCA would "want" to be in this mixture. Remember, ΔG0 is a constant that at a given T, P, and solvent conditions and depends only on the relative stability of a molecule in a given environment and not its concentration. Traube, in 1891, noticed that single chain amphiphiles tend to migrate to the surface of the water and decrease its surface tension (ST.) He observed that the decrease in ST is directly proportional to the amount of amphiphile, added up until a certain point, at which added amphiphile has no additional effect. In other words, the response of ST saturates at some point. We are more interested in what happens to amphiphiles in bulk water, not at the surface. As we showed in Figure 2 above, monomeric single chain amphiphiles are in equilibrium with single chain amphiphiles in micelles. Assume you have a way to measure monomeric single chain amphiphile in solution. What happens to its concentration as you add more and more SCA to the mixture? Turns out you observe the same effect that Traube noted with changes in surface tension. This explanation goes like this: as more amphiphile is added, more goes into bulk solution as monomers. At some point, there are enough amphiphiles added to form micelles. After this point, added amphiphiles form more micelles and no further increases in monomeric single chain amphiphiles are noted. The concentration of amphiphile at which this occurs is the critical micelle concentration (CMC). Figure $5$ shows a graph of monomeric single chain amphiphile in solution versus the concentration added to the solution. This saturation effect can be observed with other systems as well. • Consider the amount of NaCl(aq) in the solution as more NaCl(s) is added to water. At some point, the water is saturated with dissolved NaCl, and no further increase in NaCl (aq) occurs. • Consider the amount of a sparing soluble hydrocarbon (HC) in water. After saturation, phase separation occurs. Now consider the addition of a drop of a slightly soluble hydrocarbon liquid (HCL) into water, as pictured in the diagram below. At t=0, the system is not at equilibrium and some of the HC will transfer from the pure liquid to water, so at time t=0, ΔGTOT < 0. This is illustrated in Figure $6$. The following equations can be derived. \begin{array}{c} \Delta \mathrm{G}_{\mathrm{TOT}}=\left(G_{\mathrm{HC}-\mathrm{W}}\right)-\left(G_{\mathrm{HC}-\mathrm{L}}\right)=\mathrm{G}_{\mathrm{HC}-\mathrm{W}}^{0}+R T \ln [\mathrm{HC}]_{\mathrm{W}}-\left(\mathrm{G}_{\mathrm{HC}-\mathrm{L}}^{0}+R T \ln [\mathrm{HC}]_{\mathrm{L}}\right)= \ \Delta \mathrm{G}_{\mathrm{TOT}}=\left(\mathrm{G}_{\mathrm{HC}-\mathrm{W}}^{0}-\mathrm{G}_{\mathrm{HC}-\mathrm{L}}^{0}\right)+R T \ln \left([\mathrm{HC}]_{\mathrm{W}}-\ln [\mathrm{HC}]_{\mathrm{L}}\right)= \ \Delta \mathrm{G}_{\mathrm{TOT}}=\Delta \mathrm{G}^{0}+R T \ln \frac{[\mathrm{HC}]_{\mathrm{W}}}{[\mathrm{HC}]_{\mathrm{L}}} \end{array} Now add a bit more complexity to the last example. Add a hydrocarbon x, to a biphasic system of water and octanol and shake it vigorously as shown in Figure $7$. At equilibrium, x would have "partitioned" between the two mostly immiscible phases. A simple favorable reaction can be written for this system: x aq → x oct. If X is a hydrocarbon, ΔG < 0. Also, ΔGo < 0, since this term is independent of concentration and depends only on the intrinsic stability of x in water in comparison to that of octanol. This simple equation holds: \Delta \mathrm{G}_{\mathrm{TOT}}=\left(\mathrm{G}_{\mathrm{X}-\mathrm{oct}}^{0}-\mathrm{G}_{\mathrm{X}-\mathrm{w}}^{0}\right)+R T \ln \frac{[\mathrm{X}]_{\mathrm{oct}}}{[\mathrm{X}]_{\mathrm{w}}}=\Delta \mathrm{G}^{0}+R T \ln \frac{[\mathrm{X}]_{\mathrm{oct}}}{[\mathrm{X}]_{\mathrm{w}}} At equilibrium, ΔG0=0 and the equation can be rewritten as: \Delta \mathrm{G}^{0}=-R T \ln \frac{[\mathrm{X}]_{\mathrm{oct}}}{[\mathrm{X}]_{\mathrm{w}}}=-\mathrm{RTlnK}_{\mathrm{part}} where Kpart is the equilibrium partition coefficient for X in octanol and water. This can readily be determined in the lab. Just shake a separatory flask with a biphasic system of octanol and water after injecting a bit of X. Then separate the layers and determine the concentration of x in each phase. Plug these numbers into the last equation. You should be able to predict the sign and relative magnitude of ΔGo since it does not depend on concentration, but only on the intrinsic stability of the molecules in the different environments. Kpaft values are often determined for drugs since they often must diffuse across cell membranes to move into the cytoplasm where they can act. Drugs hence must have a reasonable Kpart to pass through the membrane but not so high that they are insoluble. Double Chain Amphiphiles and Bilayers In contrast to single chain amphiphiles, double-chain amphiphiles added to water form monolayers and vesicles called liposomes, as shown in Figure $8$. They can be unilamellar, consisting of a single bilayer surrounding the internal aqueous compartment, or multilamellar, consisting of multiple bilayers surrounding the enclosed aqueous solution. You can image imagine that multilamellar vesicles resemble an onion with its multiple layers. Cartoons of unilamellar and multilamellar liposomes are shown in Figure $9$, where each concentric circle represents a bilayer. Liposomes vary in diameter. They can be generally categorized into small (S, diameter < 25 nm), intermediate (I, diameter around 100 nm), and large (L, diameter from 250-1000 nm). If these vesicles are unilamellar, they are abbreviated as SUV, IUV, and LUV The chemical composition of liposomes made in the lab can be widely varied. Most contain neutral phospholipids like phosphatidylcholine phosphatidyl ethanolamine (PE), or sphingomyelin (SM), supplemented, if desired, with negatively charged phospholipids, like phosphatidyl serine (PS) and phosphatidyl glycerol (PG). In addition, single-chain amphiphiles like cholesterol (C) and detergents can be incorporated into the bilayer membrane, which modulates the fluidity and transition temperature (Tm) of the bilayer. If present in too great a concentration, single-chain amphiphiles like detergents, which form micelles, can disrupt the membrane so completely that the double-chain amphiphiles become incorporated into detergent micelles, now called mixed micelles, in a process that effectively destroys the membrane bilayer. The properties of liposomes (charge density, membrane fluidity, and permeability) are determined by the lipid composition and size of the vesicle. The desired properties will be, in turn, determined by the use of the particular liposome. The vesicles offer wonderful, simple models to study the biochemistry and biophysics of natural membranes. Membrane proteins can be incorporated into the liposome bilayer using the exact method you will be using. But apart from these purposes, liposomes can be used to encapsulate water-soluble molecules such as nucleic acids, proteins, and toxic drugs. These liposomes can be targeted to specific cells if antibodies or other molecules which will bind specifically to the target cell can be incorporated into the bilayer of the vesicle. Intraliposomal material may then be transferred into the cell either by fusion of the vesicle with the cell or by endocytosis of the vesicle. Liposomes could also be called lipid nanoparticles as they have sizes ranging up to 1000 nm. Liposomes are vesicular - with small aqueous-filled compartments. They can be made with encapsulated drugs for delivery to target cells through blood transport. They act as an emulsion in water. Lipid nanoparticles can also be particulate (insoluble), which slowly degrade and release their contents slowly in situ. Most recently, particulate lipid nanoparticles have helped save the world by being used to encapsulate messenger RNA (mRNA) for the spike protein from the SARS-CoV-2 virus which causes the COVID-19 pandemic. These lipid nanoparticles are used in the vaccine against the coronavirus mRNA molecules which encode the spike protein are "encapsulated" in the lipid nanoparticle. The mRNA contains specially modified nucleotides to increase their stability. The lipid nanoparticles also contain positively-charged lipids which help stabilize the negatively charged mRNA from degradation. The nanoparticles are endocytosed into cells, where the mRNA can be translated into SARS-CoV-2 spike protein, required for whole virus entry into the cell. The spike protein is then recognized by the immune system. The lipids used in the formulation of these nanoparticles include fatty acids, mono-, di- and triglycerols, glycerophospholipids, waxes (like cetyl palmitate), and other positively charged lipids including stearyl amine, benzalkonium chloride, cetrimide, cetyl pyridinium chloride, and dimethyldioctadecylammonium bromide. These are shown in Figure $10$ Figure $10$: Amphiphiles used to make lipid nanoparticles Why Micelles and Bilayers? Micelles and liposomes form spontaneously - i.e. ΔG < 0. But why do single chain amphiphiles form micelles and double-chain amphiphiles form bilayers? Let's think about this from a thermodynamics and structural sense. As the number of Cs in the nonpolar carbon (NC) chain increases, the ΔG for the transferring into a micelle, or by analogy, for a single chain amphiphile entering a micelle, becomes more and more negative (i.e more favored). The following equation seems to apply to the transfer of a single chain amphiphile into a micelle: ΔGo = Go (mic) - Go (aq) = + number - 709 NC \Delta \mathrm{G}^{\circ}=\mathrm{G}^{\circ} \text { (mic) }-\mathrm{G}^{\circ}(\mathrm{aq})=+\text { number }-709 \mathrm{NC} where NC is the number of carbon atoms in the chain. The first positive term depends on the nature of the head group, while the second negative term is independent of the head group. These + and - terms bring us back to the principle of opposing forces we discussed when we looked at the noncovalent interactions involved in micelle and bilayer formation. There are attractive interactions, including induced dipole-induced dipole interactions among the chains and dipole-ion, and H-bond interactions with water and the head groups. There are likewise repulsive interactions arising from steric hindrance with bulky heady groups and ion-ion repulsions. Of course, there are also entropic considerations. Let us now consider these factors as we explore what might happen to a preformed micelle as we try to put more single chain amphiphiles (sca) into it. As we increase the number of Cs in the SCA, the micelles would have a larger radius. For a given SCA with a fixed number of Cs, once a spherical micelle is formed, it can no longer retain its spherical shape if more SCAs are added. Imagine increasing the diameter of a spherical micelle 10x. A large part of the inside would have no atoms or be filled with water, which would not be favorable. Therefore, if the micelle is to grow, it can do so only by changing shape to something other than a sphere. By squeezing a tennis ball, one can imagine that the shape could distort to a circular cylinder with end caps. In this way, the acyl chains can still interact. The only problem is that head groups will now be closer than they were in the sphere. This is simplistically illustrated in Figure $11$. Imagine in a sphere the head groups radiating in a perpendicular direction from the sphere surface. As the sphere is distorted into a cylinder, the head groups would come closer together, and hence they will experience more steric interference. If a cylinder can be formed, however, it could continue to grow as long as needed with no further compression required. Imagine now that you further compressed the cylinder into a planar "bilayer" structure. The head groups would be even closer and experience even more repulsion. This bilayer will not form since growth can occur in the cylindrical phase without the added repulsion. Now consider a double-chain amphiphile (DCA). In the case of a SCA, the number (N) of head groups (HG) = the number of acyl chains (CH). Hence the surface area per HG is equal to the area per HC. Or: As/N HG = As/ N CH. For a DCA, N HG = N CH/2, therefore As/N HG = 2As/NCH. There is twice the surface area available per head group compared to that of the SCA. Therefore the DCA can tolerate more compression. It can easily be compressed to a bilayer, which as we saw, has much less As/HG. The cylindrical form has too much space per head group since water can enter the structure. The extra closeness of head groups in the bilayer can be tolerated even more since the ΔGo for transfer of a DCA into a micelle is 60% more negative than that of a SCA. The As/HG for closed vesicles differs only slightly from that of a truly planar bilayer since the vesicles are so large compared to a micelle. Once again, we have discovered that structure mediates function. We can account for the fact that SCA and DCA form micelles and bilayers, respectively, by understanding the structure of the monomers! In reality, things are more complicated. The general rule holds that single chain amphiphiles form micelles and double-chain amphiphiles form bilayers. However, under the right condition, single-chain fatty acids can form bilayers if the pH is low enough that the head group is protonated and uncharged. Why would that make a difference? Fatty acids like oleic acid would be a prime candidates for components of the membranes of protocells in the evolution of life from abiotic conditions. Likewise, short double-chain amphiphiles can make micelles. In addition, other lipids phases can be observed. Which aggregates or phases ultimately form depends on the structure of the lipid, the solvent conditions, and the temperature. These include the following phases: • lamellar gel (Lb) and lamellar liquid crystalline (La) phases • hexagonal HI (cylinders packed in the shape of a hexagon with polar heads facing out into the water • hexagonal HII (cylinders packed in the shape of a hexagon with acyl chains pointing out as in reverse micelles, and • micellar (M). We will discuss them in more detail in the next section.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/10%3A_Lipids/10.02%3A_Lipids_Aggregates_in_Water_-_Micelles_and_Liposomes.txt

Search Fundamentals of Biochemistry An overview of lipid bilayers A membrane bilayer consists of more than just two leaflets of amphiphilic leaflets. It also contains membrane proteins (which we will discuss in the next chapter) which can also be attached to carbohydrates. Most assuredly you have seen various representations of a bilayer before. Before we proceed with a more detailed description of the lipids in the bilayer and its associated properties, we present Figure \(1\) to focus our discussion. To understand the movement of lipids in an actual cell, a better understanding of lipid synthesis and trafficking in cells is important. Table \(1\) below shows the distribution of four classes of lipids in a macrophage, a type of immune cell (Andreyev, A.Y. et al) while the following figure shows how the lipids composition of membranes organelle membranes. Table \(1\): Distribution of Lipids in Resting Macrophage Lipid Categories Nucleus Mitochondria ER Plasma Memb microsome cytosol Whole cell Glycero-phospholipids 149 152 150 151 142 109 155 Prenol lipids 5 5 5 5 5 5 5 Sphingolipids 48 47 48 48 48 47 48 Sterol lipids 13 12 12 13 11 5 12 Total 215 216 215 217 206 166 220 Lipids in membranes are often distributed asymmetrically. The inner and outer leaflets of a biological membrane usually have different PL compositions. For example, in red blood cell membranes, the outer leaflet is composed mostly of sphingomyelin (SM) and PC, while the inner leaflet is composed mostly of PE and phosphatidyl serine (PS). This phospholipid contains the amino acid serine linked through its side chain (-CH2OH) to phosphate in position 3 of diacylglycerol. With a negative charge on the phosphate and carboxylate and a positive charge on the amine of PS, this phospholipid is acidic with a net negative charge. All the PS is located in the inner leaflet! This observation will become important later on when we discuss programmed cell death. A dying cell will expose PS in the outer leaflet. This is one of the markers of a dying cell. The membrane lipid composition in an average mammalian cell is shown in Table \(2\) below. Table \(2\): Membrane lipid composition in an average mammalian cell. Lipid % PC 45-55 PE 15-25 PI 10-15 PS 5-10 PA 1-2 SM 5-10 cardiolipin (bis-PG) 2-5 cholesterol 10-20 Lipid membranes also surround the variety of intracellular organelles found in eukaryotic cells. As a refresher, Figure \(2\) shows the anatomy of a typical eukaryotic cell with its variety of intracellular organelles. Figure \(3\) shows the average distribution of membrane lipids in different eukaryotic organelles. Figure \(4\), our last overview, shows the location of lipid synthesis and the resulting distribution of lipids in each leaflet. Note that most lipids are synthesized in the endoplasmic reticulum (ER). Dynamics of Membrane Bilayers Molecules are not static, but rather are dynamic. This also applies to molecular aggregates. In the first part of the section, we will discuss the rigid movement of whole lipid molecules in a bilayer, within a leaflet, and between leaflets. In the second part and the following supplement, we will consider the movement of atoms within a molecule. The movements include motions like bond bending, bond stretching, and torsion angle changes as we saw in the previous chapter section on the conformations of n-butane. The position of all atoms within a molecule can be simulated as a function of time - a molecular dynamics simulation. Such motions affect the energy of the molecule, which can be calculated for given atom positions using classical molecular mechanics and electrostatics. Liposomes and bilayers in general must be somewhat dynamic, otherwise, they would be impenetrable barriers across which nothing could pass. Cell membranes must separate the outside of a cell from the inside, but they must also allow the passage of molecules and even ions across the membrane. What is the evidence that membranes are dynamic? First, lipids can diffuse laterally in the membrane. This can be shown as follows. Make a liposome from phosphatidylethanolamine, PE, which has been labeled with TNBS (trinitrobenzensulfonate). The NH2 on the head group of PE can attach the TNBS which undergoes nucleophilic aromatic substitution with the expulsion of the SO32-. The TNB group attached to the PE head group absorbs UV light and emits light of a higher wavelength in a process called fluorescence. Next, using a fluorescent microscope, the fluorescence intensity of a region of the surface can be recorded. Then shine a laser on a small area of the surface, which can photobleach the fluorescence in the area. Over time, fluorescence can be detected from the region again. The rate at which it returns is a measure of the lateral diffusion of the labeled lipids into the region. Lipids can undergo lateral diffusion at a rate of about 2 mm/s. This implies that the lipids can transit the surface of a bacteria in 1 sec. Transverse or flip-flop diffusion (movement of a phospholipid from one leaflet to the other, not within a given leaflet) should be more difficult. Experimentally, this is investigated as shown in the diagrams below. Flip-Flop Diffusion in Liposomes: To test flip-flop diffusion in an artificial membrane, liposomes are made with a mixture of PC and a PC derivative with a nitroxide spin label (has a single unpaired electron) as shown in Figure \(5\). Both the inner and outer leaflets of the membrane have the labeled PC. Like a proton in NMR spectroscopy, a single electron has a spin that can give rise to an electron-spin resonance (ESR) signal (as a proton gives rise to a nuclear magnetic resonance signal) when irradiated with the appropriate frequency electromagnetic radiation (microwave frequency for ESR, radio frequency for NMR) in the presence of a magnetic field. The liposomes are kept at 0oC and the ESR signal is determined. Ascorbic acid, a water-soluble vitamin, and antioxidant/ reducing agent is added to the liposomes. This reduces the spin-labeled PC in the outer leaflet, but not the inner leaflet of the bilayer since ascorbic acid can not enter the liposome or otherwise interact with it. This reduces the ESR signal to a lower, constant value. The sample is divided into two. One sample is left at 0oC, the other is raised to 30oC. The ESR signal is recorded as a function of time. The 0oC prep shows no change in ESR with time, while the 30oC prep ESR signal decreases with time. This decrease results from flip-flop diffusion of labeled PC from the inner leaflet to the outer, and subsequent reduction by ascorbic acid. These experiments in experimental bilayer systems like liposomes show that flip-flop diffusion is orders of magnitude slower than lateral diffusion. Flip-Flop Diffusion in Bacterial Cells An analogous experiment can be done with bacteria. Radiolabeled 32PO4- is added to cells for one minute, which leads to the labeling of newly synthesized phospholipid (PL) which locates in the inner leaflet. The cells are then split into two samples. One sample is reacted immediately with TNBS, which will label only PE in the outer leaflet. The other sample is incubated for 3 minutes (to allow PL synthesis) and then reacted with TNBS. This is shown in Figure \(6\). After a short labeling period, the cells are destroyed by adding organic solvents which prevent new lipids biosynthesis. The lipids are extracted into the solvent and then subjected to TLC. The lipids can be labeled in three ways. Some will be labeled with 32P alone, some with TNBS alone, and some with both 32P and TNBS. TLC (or other techniques such as HPLC or GC) can easily separate PC and TNBS-labeled PC since they have different structures and hence will migrate to different places on a TLC plate. No chromatographic technique could, however, separate PC and 32P-PC, since their molecular structure is the same, the only difference being in the nuclei of the P (different number of neutrons). Those lipids with double labels (TNB and 32P) must have flipped from the inner leaflet to the outer leaflet where they could be labeled with TNBS. The cells incubated for 3 minutes before the addition of TNBS have a much higher level of doubly labeled PLs. Quantitating these data as a function of differing time of incubation at elevated temperatures show that the rate of flip-flop diffusion is much higher in cells than in liposomes, which suggests that the process is catalyzed, presumably by a protein transporter (flippase or Transbilayer amphipath transporter - TAT) in cells. The actual movement of lipids in bilayers is catalyzed by different enzymes, including flippases, floppases, and scramblases, as illustrated in Figure \(7\). Most required ATP hydrolysis for the physical movement of the lipid across leaflets. Figure \(8\) shows an interactive iCn3D model of a human plasma membrane phospholipid flippase with bound phosphatidylserine (PS) shown in spacefill. Figure \(8\): Human plasma membrane phospholipid flippase with bound phosphatidylserine (6lkn) (Copyright; author via source). Click the image for a popup or use this external link:https://structure.ncbi.nlm.nih.gov/i...kcJeKtZGDa53W6 The iCn3D model below shows the structure of a human plasma membrane phospholipid flippase with bound phosphatidylserine (PS) shown in spacefill. This protein moves PS from the outer to the inner plasma membrane leaflet, maintaining its asymmetric distribution. The other common aminophospholipid, PE, is also found predominantly in the inner leaflet. In contrast, PC and SM are found predominately in the outer leaflet. The movement of PS is from low to a high concentration and requires ATP. A build-up of PS in the outer leaflet is one signal that initiates programmed cell death (apoptosis) in the cell. Clotting is also initiated when cellular damage leads to exposed PS. Flippases are proteins that move lipids from the outer to the inner leaflet, while floppases move them from the inner to the outer leaflet. Most are also ATPases. Both promote lipid asymmetry in the membrane and floppases also help move lipids out of the cell. Scramblases moved lipids in either direction and break the asymmetry of the lipid distribution. They are important in signaling. For example, they are used to expose phosphatidylserine to the outer leaflet which promotes programmed cell death. Here are links to iCn3D models of Conformational Transitions in Bilayers If a vesicle preparation is placed in a sensitive calorimeter and the temperature slowly increased, it is observed that the vesicle preparation absorbs a significant amount of heat at a temperature characteristic of the PLs which compose the vesicle. This is analogous to what would happen if solid water was heated. At the melting point of water, an increment of heat is required, the heat of fusion, to break H-bonds and cause melting. Likewise, the heat of vaporization is measured when H-bonds are broken on the liquid-gas transition. These transitions are associated with non-covalent processes, namely, breaking H-bonds. Graphs of heat absorbed (Q) as a function of temperature, or heat absorbed/T (i.e. the heat capacity) vs temperature for the melting and evaporation of water is shown in Figure \(9\). These transitions occur at the melting point (TM) and the boiling point. The bottom heat capacity graph is nothing more than the derivative curve (or slope at each point) of the Qabs curve! Likewise, lipid vesicles undergo phase transitions comparable to the melting of water. One large phase transition at a "melting point" (TM) = 420 C can be seen in the graph of heat capacity vs temperature for vesicles made of DPPC shown in Figure \(10\). This transition is caused by conformational chains in the packing of the acyl chains of the phospholipids as the acyl chains change from trans to gauche conformations. These changes involve not the simple translation of lipid molecules within and between bilayers but rather the movement of atoms within the molecules. These kinds of motions can be modeled using molecular dynamics simulations. Before the transition, the acyl chains are more tightly packed in the gel phase, and after they are less tightly packed in the liquid crystalline phase since many chains are in the gauche conformation. A minor transition is also noted at around 360 C. This is associated with changes in the orientation of head groups. As with water going from ice to liquid, the vesicles after the phase transition are still intact. It's not like the transition of liquid to gas phase water. Vesicles in the liquid crystalline phase are more fluid, dynamic, and hence more permeable. Note that the liposomes have not been destroyed but simply have undergone a phase change, much like ice turning to liquid water. The phases of lipid vesicles are given the names gel and liquid crystalline to reflect the rigidity of the bilayer. • Gel phase (Lβ): In the gel phase, which is found at temperature < TM, the lipids are ordered with maximal packing. The acyl chains in both leaflets can be tilted so that they align in a parallel fashion (as shown in the figure below) or in a cross-tilted fashion in which they tilt toward each other. In the gel phase, the lipids diffuse slowly. This phase is sometimes called the solid phase. The gel phase is favored by low temperature and high saturation of esterified fatty acids. Saturated PC bilayers give a gel phase in the lab, • Liquid crystalline phase (Lα): In this liquid crystalline phase, which is found at temperature > TM, some saturated acyl chains have undergone all trans to gauche conformational changes. These introduce kinks into the chains which reduce packing. The notation Lα is used for bilayers of pure lipids • Liquid crystalline ordered (L0 ) and Liquid crystalline disordered (Ld): These phases typically occur with the addition of relatively high amounts of cholesterol. Cholesterol modulates the fluidity of membranes as we will see in a bit and affects bilayer properties at temperature both < and > TM. The L0 phase is often enriched in saturated (sphingo)lipids and cholesterol while the Ld phase is often enriched in unsaturated glycerophospholipids. The liquid crystalline disordered (Ld ) has fast translational diffusion and lower order while the Liquid crystalline ordered (L0) has fast diffusion with higher order. Most membrane lipids in vivo contain unsaturated fatty acids and use specific lipids for given environments to avoid the gel phase. Figure \(11\) shows some of these phases Figure \(12\) shows a snapshot of a molecular dynamics simulation of a bilayer in a gel (A) and liquid crystalline (B) phase. Note that the width of the liquid crystalline phase is smaller. Vesicles made of different PL have different TM as shown in Table \(3\) below. Table \(3\): Melting point (TM) of vesicles made with different phospholipids Lipid TM Lipid TM 12:0 PC -1 12:0 PA 31 14:0 PC 23 14:0 PA 50 16:0 PC 41 16:0 PA 67 18:0 PC 55 18:0 PA 76 18:1 PC -20 18:1 PA -8 18:2 PC -53 - - 18:3 PC -60 - - Vesicles make from phospholipids with bigger head groups have a lower TM, since they are less "stable". For example, the Tm for vesicles of di-16:0 versions of PA, PE, and PC have TMs of 67, 63, and 41 degrees C, respectively, as shown in Figure \(13\). Cholesterol and Membrane Fluidity Cholesterol is also a ubiquitous component of animal cell membranes. Its size will allow it to fit into either leaflet with its polar OH pointed to the outside. One function of cholesterol in membranes is to keep the membrane fluid at any reasonable temperature. When a membrane is at a temperature less than the TM, it is ordinarily in a gel, not a liquid crystalline phase. The cholesterol helps prevent the ordered packing of the acyl chains of the PLs, which increases their freedom of motion. Hence the fluidity and permeability of the membrane are increased. At temperatures greater than the TM, the rigid ring of cholesterol reduces the freedom of the acyl chains to rotation and hence decreases the number of chains in the gauche conformation. This decreases fluidity and permeability. Cholesterol affects membrane structure at temperatures both below and above the TM as Figure \(14\) shows the results of a molecular dynamics simulation depicting the relative order in a DMPC membrane with and without cholesterol Lipid Rafts and Nanodomains Not only are lipids asymmetrically distributed between leaflets of a bilayer, but they are also distributed asymmetrically within a single leaflet. Certain lipids often cluster within a leaflet to form lipid "rafts" which can be considered to result from lateral phase separation of the lipids within one leaflet of the bilayer. Divalent cations like calcium, which can bind to negatively charged PLs like PS, can cause "rafts" of PS to form, giving rise to lateral asymmetry within a leaflet of a bilayer. Rafts also appear to be enriched in cholesterol and lipids with saturated fatty acids, especially sphingolipids, which would lead to regions of enhanced packing and reduced fluidity. Cholesterol would stabilize packing in spaces created with lipids with large head groups. You can think of these rafts as nanodomains, analogous to the domains we observed in protein structure. Cholesterol appears to be a key player in the formation of lipid rafts. It is planar and inflexible and would pack better with saturated fatty acid chains and could also induce them to elongate to form lower energy zig-zag structures in which all the methylene groups are anti. Lipid rafts would represent a more ordered lipid phase (Lo) compared to the more disordered surrounding phase (Ld). Also compared to the structure of glycerophospholipids, the atoms in the region linking the head group and the nonpolar fatty acid chains in sphingolipids have greater potential for H bond interactions with cholesterol and other sphingolipids, as shown in Figure \(15\). Rafts probably bind or exclude the binding of other biological molecules like proteins. Some proteins are chemically modified with a glycosylphosphoinositol (GPI) group at the carboxy terminus. The PI group can insert into the membrane, anchoring the protein to the bilayer. Protein also appear to induce raft formation. Lipids rafts appear to be enriched in glycosylphosphoinositol (GPI)-anchored proteins as we will see in Chapter 11.1. Recent studies have shown that the Ebola virus interacts with lipid rafts in the process of entering and exiting the infected cell. Rafts are also involved in how cells sense and respond to their environment. Signaling molecules on the outside of the cell can bind receptor proteins in the membrane. As we will see later, conformational changes in the receptor protein signal the inside of the cells that the receptor is bound with a ligand. Once bound, the receptor can move in the membrane and often cluster in outer leaflet rafts that contain cholesterol and sphingolipids. Inner leaflet rafts have also been observed. Figure \(16\) shows two versions of an animated version of a lipid raft. The large shapes represent membrane proteins selectively found in the rafts. The most modern definition of a lipid raft is a nanoscale assembly of sphingolipids, cholesterol, and proteins that can be stabilized into platforms. Figure \(17\) shows a simplified model of a lipid raft. "The phospholipids (blue and brown) and cholesterol (yellow) are distributed in both the leaflets, whereas sphingolipids (violet) are enriched in the outer leaflet of the bilayer. The acyl chains of raft lipids are generally long and saturated (violet and brown), whereas those in non-raft domains are shorter and contain singly or multiply unsaturated acyl chains (blue). Raft domains contain concentrations of dually-acylated (green) and GPI-anchored (brown) proteins, whereas transmembrane (blue) and prenylated (green) proteins are usually non-raft associated." Lipid bilayers, in contrast to single proteins, for example, are physical mixtures. In a thermodynamic sense, entropy would disfavor raft formation as random mixing is favored. Enhance enthalpic must drive the interaction between neighboring molecules to produce nanodomains and rafts. Lipid Phase Diagrams You are familiar with the phase diagrams of water from introductory chemistry classes. Phase diagrams show the different phases that are accessible under different sets of conditions such as temperature and pressure. A traditional phase diagram for water is shown in Figure \(18\). The horizontal dotted line at 101 kPa shows the states of water as a function of temperature at 101 kPa = 1 atm pressure. The phase transition of solid to liquid water occurs at 00 C (freezing/melting point of water) while the liquid to gas transition occurs a 1000 C (boiling point of water). At a reduced pressure (0.6 atm), all three phases of water can exist at 0.01 0 C, the triple point of water. In an analogous fashion, lipid bilayers have phase transition diagrams as well. Instead of showing phases as a function of temperature and pressure, they are usually shown as a function of temperature and concentration of a specific lipid component such as cholesterol, which as described above affects TM, fluidity, and raft formation. An example of a theoretical phase diagram for membranes composed of saturated dipalmitoylphosphatidylcholine (16:0), the most common saturated fatty acid in animals, plants, and microorganisms vs cholesterol content is shown in Figure \(19\). DPPC vesicles have a TM = 41 0C or 314 K. The phase diagrams show that when cholesterol is added to DPPC bilayers at temperatures above TM, the bilayers change from the liquid-disordered (Ld) phase to the uniform liquid-ordered (Lo) phase at around 20 mol% cholesterol. At cholesterol concentrations between around 10-20 mol% and temperatures just above the TM, both the L0 and Ld phases coexist. The coexistence of two phases mimics a raft. At low cholesterol levels, the bilayer changes from the gel (or Lβ) to the Ld phase. At really high cholesterol, only the L0 phase exists. This makes sense as the fatty acids are all saturated and cholesterol's rigid rings reduce the freedom of the acyl chains to rotation and hence decrease the number of chains in the gauche conformation. Yet as an "impurity" (cholesterol) has been added to the system, the system is less rigid, and more fluid-like. From 0-7 mol% cholesterol, two phases exist, the gel (or Lβ) and the Ld phase. Between 7-23 mol% cholesterol, a combination of phases is seen. As mentioned above, the outer leaflet of mammalian plasma membranes is composed mostly of sphingomyelin (SM) and PC, while the inner leaflet is composed mostly of phosphatidyl ethanolamine (PE) and phosphatidyl serine (PS), along with cholesterol. These don't appear to separate into Ld and L0 phases and appear not to form nanodomains or rafts. In the lab, outer membrane lipids easily form vesicles but the inner leaflet polyunsaturated PEs form other phases (hexagonal or cubic). It appears that asymmetric lipid distribution is critical for biological bilayer formation. Since nanodomains are difficult to observe and study experimentally, molecular dynamics simulations are used to provide insight into their structure and properties. The right-hand images to the right show snapshots of the membrane from molecular dynamic simulations. Cholesterol is shown in white in Figure \(19\). 8 shows the membrane in the Lo phase, 5 shows it in the Ld phase (note the reduced membrane width), and 3 in an ordered (blue box) and disordered (red box) phase. The simulations support the theoretical phase diagram showing the coexistence of the Ld and L0 phases as well as finding a hexagonal-closest packed cholesterol poor with Ld domains in the region of the phase diagram showing the coexistence of both Ld and L0 phases. In addition, cholesterol is excluded from most ordered regions. This is shown in Figure \(20\) showing a top-down view of the membrane. The dark green lipid headgroups are those that are hexagonally closest packed, an ideal you will remember from introductory chemistry courses. Now imagine what a phase diagram would look like for a three (DOPC, sphingomyelin, and cholesterol) or more component system! We won't show any but from the "simple" two-component system described above, it should be evident that we have a long way to go before understanding the complexity of membrane bilayers. Ternary bilayers system often from more macroscopic (vs nanoscopic) domains (or rafts) which can be studied using fluorescence microscopy. The actual biological membrane must be able to adopt very nonplanar shapes with positive and negative curvature. Membranes must also be able to fuse (for example the egg and sperm). Another lipid phase, which we have not yet discussed, may be involved. New lipid phases of a single membrane lipid can form based on the relative percentages of lipid and water. These include hexagonal phases. Figure \(21\) shows an image of a hexagonal phase of phosphatidylethanolamine with 16:0 fatty acids. Note the water inside of the middle ring of PE molecules. Figure \(22\) shows an interactive iCn3D model that shows the inner ring outlined in the red circle. Note that water surrounds each of the closest packed lipid tubules which extend back into the figure in 3D. This phase creates an aqueous channel through the interior of each tubule. Figure \(22\): Hexagonal phase of phosphatidylethanolamine with 16:0 fatty acids (Copyright; author via source). Click the image for a popup or use this external link: not available Figure \(23\) shows variants of the hexagonal phase as well as a cubic phase of lipids. Double-chain amphiphiles with a small polar head (like PE and PA) are more likely to form a hexagonal II phase with elongated tubules than are more cylindrical lipids like PC. Note that if a single fatty acid is removed from a double-chain amphiphile like PE, a narrow conical shape single-chain amphiphile arises, which can either form a micelle (not shown) or a hexagonal I phase. In vitro, equimolar amounts of PC (which forms the laminar phase) and PE (which can form an HII phase) can either form the HII phase(at low aqueous pressure) or a lamellar phase (at high aqueous pressure). Are hexagonal lipid phases found in biological membranes? The answer is likely yes in the formation of unusual cellular structures. The plant plasmodesmata, shown in Figure \(24\), is one such structure. Just focus on the two bilayers connected by the membrane-lined membranes of the channel. Figure \(24\): Plant plasmademata. Yanbiao Sun et al. https://www.nature.com/articles/s41438-019-0129-3. Creative Commons license. http://creativecommons.org/licenses/by/4.0/. Plasmodesmata are membrane-lined channels that cut across the plant cell wall and directly connect cells, allowing the flow of water and nutrients between the cells. HII phases have been seen in the endoplasmic reticulum membrane. These membranes, along with the mitochondrial inner membrane and the inner membrane of chloroplasts are highly curved and contain higher concentrations of lipids that allow that curvature as well as the formation of H II phases. The function of some membrane enzymes as well as processes such as membrane fusion and fission are enhanced by HII-forming lipids. Piecing it all together Our emerging understanding of lipid structure has taken us from micelles and vesicles to the complexity of actual biological membranes with different phases and nanodomain structures. This complexity is needed as membranes must be dynamic in ways the proteins, for example, aren't. They must be able to pinch off either as extracellular or intracellular vesicles, for example, in the process of exocytosis and endocytosis. Both positive and negative curvatures of the membrane must be enabled. The incredible complexity of the "Lego-like" lipid monomers that assemble and rearrange into every fluctuating membrane is yet another exquisite illustration of our repeat mantra, that structure and shape mediate all function. The "Lego-like" membrane monomers in various phases and regions of membrane curvature are illustrated in Figure \(25\). Given their shape, some glycerolipids don't even appear to spontaneously form bilayers by themselves. The shape and type of fatty acids found in the membrane lipids will determine the local properties of the membrane, its phases, and the presence of nanodomains. The activity of membrane enzymes, and membrane fission and fusion events, will also depend on the local properties of membranes. The figure above. doesn't even account for the presence of peripheral and integral membrane proteins, which we will discuss in the next section. As we learned with proteins, we can be misled by looking at beautiful but static images of protein. Their dynamic motion is critical to their function. In addition to the dynamic membrane events discussed above, there is a myriad of other events that take place at and in membranes. Here are a few. Other dynamic events in bilayers Membranes are not static. They are synthesized, their contents shuffled, they fuse with other membranes, and large proteins are inserted into them. Let's explore some of these. Membrane Trafficking Movement of key "cargo" molecules into (endocytosis) and out of (exocytosis) the cells occurs mostly through membrane-encapsulated vesicles. Vesicles contain all types of biological molecules including lipids, both synthetic and absorbed. Part of the differences in lipid composition between membrane layers and between different organelles derived from this highly orchestrated and controlled movement of vesicles. Details are shown in Figure \(26\)below. In eukaryotes, the biosynthetic secretory pathways move molecules from the endoplasmic reticulum to the cis Golgi (CGN) to the trans-Golgi (TGN) and to the plasma membrane (for integral membrane proteins) or for secretion. Since most lipids are synthesized in the endoplasmic reticulum (ER), their distribution to different locations in cells is critical in maintaining the asymmetric distribution of lipids found in cells. Fusion of membranes: Fusion Peptides Another dynamic event in membranes is the fusion of two bilayers from two different cells vesicles or of a vesicle and cell membrane. These events are facilitated by fusion peptides. Figure (27\) below shows a molecular dynamics simulation snapshot showing how a fusion peptide in a single DMPC bilayer causes a constriction of the bilayer with the two leaflets approaching each other. Figure (28\) below shows the area per lipid (APL). Insertion of Membrane Proteins Another dynamic event is the insertion of a membrane protein. Now let's look at changes in the bilayer on insertion of the voltage-dependent anion channel (VDAC) membrane protein, as shown in Figure (29\) below. Panel a shows the voltage-dependent anion channel (VDAC) is shown in a cartoon β-barrel representation. Residue E73 and the water molecules nearby are represented as spheres. Serine and threonine residues constituting hydrophilic areas close to E73 are shown in ball and stick representation. In the snapshot, a DMPC lipid is shown flipping close to the E73 and K110 residues. Panel b shows top view perspectives of the circular representation of the local thickness, calculated considering phosphorus atoms, and area per lipid. Introduction to Lipid Signaling - Chemical Cleavage of Membrane Lipids Everything in this chapter so far describes the structure and dynamics of the components of the lipid components of a bilayer. The dynamic changes described involve the physical movement of lipids molecules in the membrane. Let's briefly introduce another type of movement that involves not only physical but chemical changes in the cell. What happens when specific lipids in membranes are chemically cleaved by lipases, enzymes which are analogous to protease? It turns out these changes lead to signaling within the cell. We will only briefly introduce lipid signaling in this chapter, but explore it in more detail in Chapter 12. Lipids are not just used as a passive component of membranes, or as a source of stored energy. They are involved in the process of signal transduction at the cell membrane, a process by which the interior components of the cell respond to a signal external to the cell, allowing the cell to respond to its local environment. Usually, a chemical signal on the outside of the cell is the "primary messenger" that causes the cell to respond. Usually, the chemical transmitter of information does not get into the cell. Rather it binds to surface receptors on the cell membrane surface. Somehow, the cells sense that a ligand is bound to the outside. Enzymes, usually in the membrane or at the intracellular surface of the lipid bilayer are activated. Many of these enzymes cleave lipids in the membrane. The cleaved fragments of the lipid molecules serve as intracellular signals or "secondary messengers", which can bind to intracellular enzymes to activate intracellular processes. Figure (30\) below shows some of the lipid mediators which are generated by the process and signal the cell to respond. Figure (30\): Membrane lipids involved in signaling Fatty acid amides are potent mediators of neurological processes. In one interesting experiment, sheep were sleep deprived. Reasoning that the brain might release a biochemical signal into cerebrospinal fluid to induce sleep, scientists at Scripps removed some of this fluid and isolated a substance that was not found in rested sheep. On analysis, the structure was shown to be an amide of oleic acid. Oleylethanolamide has been shown to bind to the peroxisome-proliferator-activated receptor-a (PPAR-a) which resides in the nucleus. This ligand, by affecting gene transcription, appears to regulate body weight and the feeling of fullness after eating (satiety) as it leads to reduced eating. People have sought the natural neurotransmitter which binds to the same receptor in the brain as THC, the active ingredient of marijuana. The endogenous cannabinoid is an amide of arachidonic acid, anandamide. Figure (31\) below shows the structures of key fatty acid amides and THC. Figure (31\): Fatty Acid Amides: Neurochemical Mediators This fatty acid amide is an example of a class of lipid derivatives called N-acylethanolamines (NAEs). These molecules, with acyl groups that vary in the number of carbons and double bonds, are found widely in organisms in nature. Naturally occurring anandamide leads to increased food intake after a short period of reduced food intake. One of the known physiological effects of THC is increased food consumption (the munchies). Lucanic et al (2011) have shown that decreases in NAEs extend the life span of the small roundworm C. elegans, which has become a model organism to study genes in eukaryotes. Caloric restriction has been shown to increase the life span in a variety of organisms. In invertebrates, anandamide seems to inhibit food intake, even in organisms that lack a receptor similar to which cannabinoids bind. This might seem paradoxical in that anandamide (and THC) in humans seems to induce eating. However, under long periods of caloric restriction (low-level starvation) in rats, anandamide levels are suppressed, leading to a low energy-consuming state. In general, it appears that reductions in NAEs occur during periods of caloric restriction. Mutant worms which have reduced levels of NAEs through targeted enzyme disruptions that affected either NAE synthesis or degradation have longer life spans. If normal (wild-type) worms were placed under caloric restriction but given EPA-ethanolamine (the most abundant NAE in these worms), they did not have an extended lifespan. Membrane Monolayers - Lipid droplets and lipoproteins We explored membrane bilayers that contain two leaflets above. These bilayers separate the outside and inside of cells as well as the outside and inside of internal organelles. It turns out that there are two main types of lipid-encapsulated structures in which only one phospholipid leaflet separates the interior contents from the outside. These are lipid droplets and lipoproteins. In both cases, the monolayers encapsulate TAGs and cholesterol esters - hydrophobic molecules, so there is no need for another inner leaflet to stabilize an encapsulated aqueous environment. Lipid Droplet formation So how are triacylglycerols stored in cells? In lipid droplets! In contrast to the common single and double-chain amphiphiles, which have charged atoms in their head groups and which form micelles and bilayers, respectively, triacylglycerols (TAGs) and cholesterol esters (CEs), which are almost completely nonpolar, coalesce into lipid droplets in cells. These droplets can range from very big, which are found in adipocytes (fat cells) where they take up almost all of the available space and where they are used for energy storage, to small, which are found in all cells, where they are used mostly for membrane biogenesis and energy mobilization. When esterified into esters, fatty acids, and steroids also pose less potential toxicity to cells. Lipid droplets are often found in close approximation or attachment to mitochondria, endoplasmic reticulum (ER), and peroxisomes (where plasmalogens with ether-linked fatty acid instead of ester-linked are synthesized), all organelles intimately involved in membrane and energy biochemistry. Many of the enzymes (acyltransferases for example) required for TAG metabolism are found in the mitochondria and the ER. Since lipids droplets are specialized to cells and don't form if TAGs are just added to water, we'll discuss their structures and their dynamic assembly in this section. The droplets are now considered actual cellular organelles. In contrast to other organelles which are bounded by a bilayer member, lipid droplets are surrounded by a monolayer of phospholipids which prevents exposure of the nonpolar contents to the aqueous cytoplasm. PC and PE appear to be the major phospholipids in the monolayer and both are synthesized mostly in the ER. There are many different proteins involved in the formation and interaction with lipid drops, including • perilipins: There are multiple types of perilipins. Perilipin 1 is found in adipocytes and cells that synthesize steroids (adrenals, ovaries, and testes). Perilipin 2 and 3 are found in most cells • Acyl CoA synthetases and acyltransferase: These enzymes activate free fatty acids for metabolic processes. • seipins: These are involved in lipid droplet shape number, and size. It appears to be involved in the formation of lipid drops and moving them from the ER to the cytoplasm. Facilitates initiation of LD formation, and ensures that vectorial budding of LDs from the ER is directed toward the cytoplasm It appears that lipid droplets arise from the ER which are involved in membrane biogenesis and "trafficking" of membranes to different locations in the cell. A general structure of a lipid droplet is shown in Figure \(32\). The left figure shows just internal triacylglycerols (TAGs), but the inside would also contain cholesterol ester (fatty acid esterified to the cholesterol OH) and the monolayer would also contain unesterified cholesterol. (32\): Cartoon view of a lipid droplet. How might these complex lipid droplets form in a cell if not by phase separation? With the help of proteins. of course. Figure \(31\) shows how newly synthesized TAGs (made in the ER membrane, could self-aggregate in the bilayer to form a "lens" which on further growth and addition of lipid binding proteins could bud off into the cytoplasm to form the droplets. Figure \(33\)s shows an incredible image of diacylglycerols (DAG) accumulating in a bilayer to form a clear lens in the membrane. This was produced by a molecular dynamics simulation. Figure (34\) below shows a cartoon illustrating the synthesis of a lipid droplet from the ER membrane. A membrane protein not displayed in the above figure, seipin, denotes the location for lipid droplet formation in the ER membrane. Mutants of the protein are associated with lipodystrophy. In yeast, seipin and another membrane protein Ldb16, associate to allow lipid droplet formation. Seipin aggregates to form a homo 10-mer in the membrane but in contrast to human cells, it alone can not concentrate triacylglycerol in the membrane. It requires the binding of another protein, Ldb16, for that to happen. Figure (35\) below shows yeast, human and fly seipin and their properties. Figure (35\): Yeast, human and fly seipin and their properties. Nat Commun 12, 5892 (2021). https://doi.org/10.1038/s41467-021-26162-6. Klug, Y.A., Deme, J.C., Corey, R.A. et al. Mechanism of lipid droplet formation by the yeast Sei1/Ldb16 Seipin complex. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/. Panel A shows a lipophilicity potential on the surface of yeast (left), human (middle), and fly (right) as viewed as (i) homodecamer assembly from the cytosol, (ii) individual protomers, or (iii) transparent overlay over zoomed in cartoon representation of the central α1-α2 helices. Surfaces are colored from hydrophilic (dark cyan) to hydrophobic (gold). Panel B shows a side view of the luminal domains of yeast, human (PDB 6DS5) and fly (PDB 6MLU) Seipin in relation to the plane of the ER membrane (indicated by a dotted line). Panel C shows the charge distribution of the yeast Sei1 central helices (α1, α2), depicted as a transparent Coulombic electrostatic potential surface representation (Red, negative charge; blue, positive charge; white, no charge) overlayed over a cartoon representation (light blue) to show acidic side chains. Panel D shows a top view of molecular dynamic simulations of Sei1 in a POPC membrane with 3% trioleylglycerol. Images depict the average lipid number density of trioleylglycerol. Inset – zoom of the corresponding box showing positions of TM1 and TM2. The lumenal domains form the ring with a floor as shown in Panel A above. In addition, the transmembrane segments for the cage top and sides. A switch area between the lumenal and transmembrane segments which can occupy two different conformations appears important for function. The closed cage allows accumulation and hence phase-separation of triacylglycerols, while the open form allows the nascent droplet to grow and then bud. The Ldb16 has helical regions enriched in serine and threonine, as is required for TAG loading. These -OH-containing amino acids are present in seipins in humans and flies. Site-specific mutations of the serine and threonines in the region of Ldb16 lead to problems with lipid droplet formation. Figure (36\) below shows a cartoon of the serine- and threonine-enriched helix in Ldb16. Figure (36\): Cartoon of Ldb16 predicted helix enriched in serine and threonine. Klug et al, ibid. Another model for the assembly of lipid droplets by the Sei1-Ldb16 yeast complex is shown in Figure (37\) below. Figure (37\): Sequential TAG interactions mediate LD assembly by the Sei1-Ldb16 complex. In the ER bilayer, TAG molecules (blue) concentrate in the proximity of Seipin oligomers (orange) via weak interaction with Sei1 TMs. TAG molecules within the ring interact strongly with Ldb16 (green) hydroxyl-containing residues, facilitating TAG coalescence and lens formation. Klug et al, ibid. Figure \(38\) shows an interactive iCn3D model of the homo 10-mer yeast seipin membrane complex (7OXP). Figure \(38\): Yeast seipinhomo 10-mer yeast seipin membrane complex (7OXP). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...wMLHBncjFZMbFA The two transmembrane segments (helices) from each of the monomers are evident by rotating the structure so the homo 10-mer ring is viewed from the side. Lipoproteins We eat, digest, and transport dietary fat. We also make fat and transport it through the blood as well. We saw that free fatty acids are carried in the blood by the most abundant protein in the blood, albumin. What about the very insoluble triacylglycerols and cholesterol esters? Turns out they are also transported in the blood by nanoparticles similar to lipid droplets. They are called lipoproteins since, like lipid droplets, they have proteins associated with them. Lipoproteins vary in density and size. The densest is called high-density lipoproteins (HDL). As they get larger and more filled with, they form less dense lipoproteins (low density - LDL, intermediate density - IDL, and very low density - VLDL). These contain nondietary lipids made by organs like the liver. Dietary fats are processed in the intestine into very large particles called chylomicrons. Their relative size and density are shown in Figure \(39\). Introduction to Lipids and Lipoproteins. Kenneth R. Feingold, MD. Creative Commons (CC-BY-NC-ND) license. A copy of the license can be viewed at http://creativecommons.org/licenses/by-nc-nd/2.0/. With permission. As with lipid droplets, lipoproteins have a single outer monolayer leaflet containing double chain membrane lipids like phosphatidylcholine and free cholesterol Inside are the triacylglycerols and cholesterol esters. Proteins are bound to the outer monolayer. Figure \(40\) shows two renderings of discoidal HDL particles containing a single type of protein, Apo-A1. The TAGs are shown in cyan line rendering on the inside, along with cholesterol esters (in spacefill). The bottom part of the figure shows the polar Ns and Os decorating the outer part of the monolayer of phosphatidylcholine surrounding the TAGs and cholesterol esters. Figure \(41\) shows an interactive iCn3D model of discoidal HDL (3k2s) Figure \(41\): discoidal HDL (3k2s) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...BE5XYUopbjGALA Table \(1\) below shows the proteins associated with the different types of lipoproteins. Lipoprotein Density (g/ml) Size (nm) Major Lipids Major Apoproteins Chylomicrons <0.930 75-1200 Triglycerides Apo B-48, Apo C, Apo E, Apo A-I, A-II, A-IV Chylomicron Remnants 0.930- 1.006 30-80 Triglycerides Cholesterol Apo B-48, Apo E VLDL 0.930- 1.006 30-80 Triglycerides Apo B-100, Apo E, Apo C IDL 1.006- 1.019 25-35 Triglycerides Cholesterol Apo B-100, Apo E, Apo C LDL 1.019- 1.063 18- 25 Cholesterol Apo B-100 HDL 1.063- 1.210 5- 12 Cholesterol Phospholipids Apo A-I, Apo A-II, Apo C, Apo E Lp (a) 1.055- 1.085 ~30 Cholesterol Apo B-100, Apo (a) Table \(1\): Proteins associated with the different types of lipoproteins.Introduction to Lipids and Lipoproteins. Kenneth R. Feingold, MD. Creative Commons (CC-BY-NC-ND) license. A copy of the license can be viewed at http://creativecommons.org/licenses/by-nc-nd/2.0/.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/10%3A_Lipids/10.03%3A_Membrane_Bilayer_and_Monolayer_Assemblies_-_Structures_and_Dynamics.txt

Search Fundamentals of Biochemistry Introduction Lipids, although small compared to large biopolymers like proteins, nucleic acid, and large glycans, are very heterogenous in structure, given the large array of fatty acid and isoprenoid chain lengths, numbers of double bonds, etc that appear in different lipid classes. In addition to analyzing lipid structure, lipids are used in the laboratory to create liposomes (vesicles), which serve as models for membrane bilayers and encapsulation of chemical species (drugs, vaccines) for medical use and solubilization of membrane proteins. In this section, we will concentrate on the creation of lipid vesicles (critical for the encapsulation of RNA vaccines against the SARS CoV-2 spike protein) and the chemical analysis of biological lipids, whose composition affects health and disease states. Liposomes Liposomes produced in the lab can be unilamellar, consisting of a single bilayer surrounding the internal aqueous compartment, or multilamellar, consisting of multiple bilayers surrounding the enclosed aqueous solution. You can imagine the multilamellar vesicles resemble an onion with its multiple layers. Cartoons of unilamellar and multilamellar liposomes are shown in Figure \(1\), where each concentric circle represents a bilayer. Liposomes vary in diameter. They can be generally categorized into small (S, diameter < 25 nm), intermediate (I, diameter around 100 nm), and large (L, diameter from 250-1000 nm). If these vesicles are unilamellar, they are abbreviated as SUV, IUV, and LUV, respectively. Their various sizes are shown in Table \(1\) below, in comparison to other large biological structures. Table \(1\): Sizes of liposomes/vesicles compared to other biological structures The chemical composition of liposomes can be widely varied. Most contain neutral phospholipids like phosphatidylcholine (PC), phosphatidyl ethanolamine (PE), or sphingomyelin (SM), supplemented, if desired, with negatively charged phospholipids, like phosphatidyl serine (PS) and phosphatidyl glycerol (PG). In addition, single-chain amphiphiles like cholesterol (C) and detergents can be incorporated into the bilayer membrane, which modulates the fluidity and transition temperature (Tm) of the bilayer. If present in too great a concentration, single-chain amphiphiles like detergents, which form micelles, can disrupt the membrane so completely that the double-chain amphiphiles become incorporated into detergent micelles, now called mixed micelles, in a process that effectively destroys the membrane bilayer. Given the large degree of unsaturation at C2, what do you expect the transition temperature of a liposome composed only of egg yolk PC to be? (Vesicles made using more saturated PC from mammalian sources have Tm of around 40oC.) This high degree of unsaturation makes egg yolk PC very susceptible to oxidation, which could alter the properties of the liposome dramatically. Synthetic PC made with saturated fatty acids could alleviate that problem. The properties of liposomes (charge density, membrane fluidity, and permeability) are determined by the lipid composition and size of the vesicle. The desired properties will be, in turn, determined by the use of the particular liposome. The vesicles offer wonderful, simple models to study the biochemistry and biophysics of natural membranes. Membrane proteins can be incorporated into the liposome bilayer using the exact method you will be using. But apart from these purposes, liposomes can be used to encapsulate water-soluble molecules such as nucleic acids, proteins, and toxic drugs. These liposomes can be targeted to specific cells if antibodies or other molecules which will bind specifically to the target cell can be incorporated into the bilayer of the vesicle. Intraliposomal material may then be transferred into the cell either by fusion of the vesicle with the cell or by endocytosis of the vesicle. Since phospholipids will spontaneously form some type of bilayer structure when placed in water, most efforts in liposome production involve producing vesicles with the desired size, lamellar structure, and physical characteristics, which as previously stated are controlled both by liposome size and chemical composition. Also, ways must be developed to entrap the desired molecule inside the vesicle in the most cost-effective manner, and with minimal leaking of contents. All methods of production involve drying of organic solvent-solubilized lipids, dispersion of the lipids in the appropriate aqueous solution, and formation of monolamellar (one bilayer) liposomes or vesicles. Finally, the vesicles are characterized (chemical composition, Tm, permeability, size, etc.) Drying of lipids Purified lipids of the desired composition (often egg PC:cholesterol:PS in molar ratios of 0.9:1.0:0.1) are dissolved in a purified, water-free organic solvent mixture (often chloroform/methanol, 2:1 v/v) and dried down in a round bottom flask on a rotary evaporator under reduced pressure (using a water aspirator) and slightly elevated temperature (20-40oC). The rapid rotation of the flask will ensure that the lipid is dispersed over a large surface area, and will increase the rate of evaporation. To remove the last traces of solvent, the dried flask is usually placed under a high vacuum overnight. If a small volume (< 1 ml) of lipid solution is used, the solvent can be evaporated under a stream of nitrogen. To avoid entrapment of residual chloroform in the lipid film, the film is dissolved in t-butyl-methyl ether or diethylether, and dried several times. Alternatively, the residual solvent can be removed under a high vacuum. Dispersion of the lipids There are three main methods of dispersing lipids into an aqueous solution to form liposomes. a. mechanical dispersion - in this method, lipid dried onto the inside glass surface of a container is hydrated with an aqueous solution, which peals off the lipid to form multilamellar - MLV - (multiple bilayers separated by water) vesicles. Only a small part of the aqueous solution is encapsulated inside the liposome, so this is not the method of choice for the encapsulation of expensive or rather insoluble solutes. Depending on the degree of agitation and the nature of the lipid used, different-sized liposomes can be prepared. b. organic solvent dispersion - In these methods, the lipids, which are dissolved in organic solvents, are injected through a fine needle, at a slow rate, into an aqueous solution in which the organic solvent may be miscible (such as ethanol) or immiscible (such as ether). In each case, the lipids orient at the interface between the organic solvent and aqueous solution, to form bilayer structures. Injection of ethanol-dissolved lipids provides a simple way to produce SUV, but because liposome formation can not occur at an ethanol concentration greater than 7.5%, only a fraction of the total aqueous phase can be entrapped in the vesicle; hence this technique is not cost-effective for entrapment of an expensive solute. Alternatively, the lipid can be dissolved in ether and slowly injected into an aqueous solution which is warmed so that the ether evaporates at the rate at which it is injected. Since the ether is volatilized, large amounts of lipid can be introduced and the encapsulation efficiency of the aqueous solution is high. c. detergent dispersion and solubilization - In this method, lipids are solubilized in an aqueous solution through the addition of detergents. The detergents are removed slowly from the solution, resulting in the spontaneous formation of liposomes. Detergents are single-chain amphiphiles that spontaneously form micelles in an aqueous solution when the concentration of free lipid rises to a minimum critical value, the critical micelle concentration (CMC); at this concentration, self-association of detergent results in the formation of a stable aggregate, the micelle. This is illustrated in Figure \(2\). Table \(2\) below shows the properties and CMC of various detergents. Table \(2\): Properties and CMC of various detergents (data from Avanti Polar Lipids) Name mM mg/ml MW n-hepty glucopyranoside 70 19.5 278 n-octyl glucopyranoside 23.2 6.8 292 n-nonyl glucopyranoside 6.5 2.0 306 n-decyl maltoside 2.19 1.1 499 n-dodecyl maltotrioside 0.2 0.16 825 Triton X-100 (a) 0.24 0.15 625 Nonidet P-40 (b) 0.29 0.02 603 Tween 20 (c) 0.033 0.04 1364 Brij 98 (d) 0.025 0.04 1527 sodium deoxycholate 2-6 1.7 415 sodium taurocholate 10-15 6.7 538 sodium cholate 14 6.0 431 sodium dodecyl sulfate 8.3 2.4 289 Making liposomes by dialysis Lipids are deposited in a small container. An aqueous solution is then added, containing water-soluble molecules for encapsulation. Detergent is then added at a concentration in excess of the lipid concentration and greater than its CMC. The lipid molecules are then "emulsified" in the detergent micelle. The solubilized mixture is then placed in a semi-permeable dialysis bag, which is placed in a large volume of an aqueous solution. The free detergent in solution is in equilibrium with the detergent in the micelle. The bag contains microscopic holes large enough for the monomeric detergent molecule to pass through, but small enough so that the large micelle can not. The lipid, during this process, is embedded in the micelle forming a detergent-lipid mixed micelle. As dialysis continues, the monomeric detergent partitions throughout both the volume in the bag and the volume surrounding the bag, while the mixed micelle remains in the bag. If the aqueous solution surrounding the bag is changed several times with fresh solution, the equilibrium in the bag is shifted to the monomeric form. Alternatively, detergent-adsorbing beads (such as Bio Bead SM-2 by Bio-Rad) can be placed in the aqueous solution surrounding the bag to speed up the process of detergent re-equilibration. Eventually, all the detergent is in this form, and during the process, which occurs slowly, the lipid in the mixed micelle self-associates to form a liposome. A detergent of low monomer molecular weight and a high CMC is most desirable for this method of liposome production. Another method of removing the free detergent is through gel filtration chromatography. In this technique, molecules of disparate molecular weights can be separated from each other. An explanation follows this discussion. This method of formation of unilamellar liposomes is the method of choice if membrane proteins are to be inserted into the liposome bilayer to target the liposome. It is not the best method, however, for quantitative encapsulation of expensive soluble molecules. Making Liposomes by Extrusion These multilamellar liposomes can be further processed to form unilamellar liposomes by several techniques. These include probe or bath sonication of the MLVs, extrusion at high pressure of the MLVs through membrane filters of defined pore size, or pH-induced vesiculation in which a transient change in pH destabilizes the MLVs in favor of unilamellar liposomes. Another technique involves the fusion of SUVs by repeated freezing and thawing or by fusion of SUVs containing acidic phospholipids (such as PS) through Ca2+-mediated aggregation. Figure \(3\): below shows the structure of vesicles as they undergo multiple freeze/thaw cycles. Figure \(4\) shows the final step in making large unilamellar vesicles by extrusion of freeze/thaw intermediates. Once the liposomes are formed, they must be separated from free monomeric lipids, detergents, and unencapsulated solutes. This can be done again by dialysis, or more readily by size exclusion chromatography. Macromolecules of different sizes can be separated on a column in which the stationary phase is a polymerized agarose or acrylamide bead, which contains pores of various sizes. A small molecule (such as monomer detergent, free lipid, or small aqueous solute) in the mobile phase (aqueous buffered solution) may enter the pores in the bead, while a larger macromolecule or aggregate (such as a large protein, a micelle, or a liposome) may not, due to size restriction. The result is that a larger fraction of the overall volume of the column is available to the smaller molecules, which thus spend a longer time on the column and are eluted by the mobile solvent after the larger species. Liposomes can be characterized both chemically, to determine the average lipid and protein (if they were incorporated) composition of the bilayer, and physically, to determine the size, permeability, lamellarity, and amount of encapsulated material. Size is usually determined by electron microscopy or indirectly by light scattering from these large species. Chemical Analysis of Lipids There are so many lipids and their derivatives that are so subtlety different that the most sensitive ones are needed for separation and analysis. The field of lipidomics focuses on the analysis of the structure and function of all lipids in the cell. The most useful techniques are analysis (and separation) by gas chromatography (GC) followed by mass spectrometry (MS). NMR spectroscopy is also important. For GC analysis, the lipid must be made volatile, which limits its use in some circumstances. MS analysis is the most sensitive. MS requires ion formation, and techniques like electrospray ionization and matrix-assisted laser desorption/ionization (MALDI) are used. Gas chromatography (This section is adapted from https://www.intechopen.com/chapters/64008. Creative Commons Attribution 3.0 License). In GC, gas is the mobile phase that carries lipids through the stationary phase of the column. Gas chromatography (GC) is used to separate organic compounds from a mixture in the gas form. For this purpose, the GC uses interactions among the sample components and the stationary/mobile phases. After lipid extraction with chloroform and/or methanol, the samples (lipid mixture) are usually liquids and must be exposed to a high temperature at the gas chromatograph entrance (injector). Vaporized, the samples are carried by a gas, which is usually a nonheavy and inert gas (i.e., hydrogen, helium), through a long capillary column containing a high or low polarity material (stationary phase) The gaseous compounds generated from the vaporized sample interact with the stationary phase which allows each compound to elute/separate at a different time (retention time). Because GC considers both the chemical and physical properties of the vaporized compounds, those with more chemical affinity to the stationary phase will take a longer time to be removed from the column and the temperature will influence the overall process. This explains why the column stays in an oven, which is programmed to work at different temperature ranges (i.e., temperature programming) in which the compounds are carried out by the gas according to their boiling point until they get to an electronic detector. At the end of GC analysis, the electronic detector generates a chromatogram based on retention time by intensity. This allows a qualitative identification of the lipid compounds by comparing their retention times with certified standards using the flame ionization detector (FID) or by deduction of spectra information using a mass spectrometer as a detector. Lipid quantification can also be performed using analytical procedures of external or internal certified standards in GC analysis. The main points to be considered when assessing FAs by GC analysis are the carrier gas flow rate, column length, and temperature because these can influence the order or retention time of the lipid compounds and then must be precisely standardized. The column length of the stationary phase influences the resolution of the analytes, once the number of theoretical plates (hypothetical zone in which two phases establish an equilibrium with each other) is respectively high in the longer column. As fat and oils have high boiling points not supported by the stationary phase, a previous derivatization reaction step is required after lipid extraction from the biological sample, in which triacylglycerol and free fatty acids are transformed into their respective free fatty esters with lower boiling points (transesterification/esterification reaction). Several methods are available for FAs derivatization. Particularly for cholesterol analysis, the sample preparation must consider a derivatization reaction. This allows to block protic sites of steroids obtained after an unsaponifiable lipid extraction had been performed, and also, diminishes dipole-dipole interactions, increases the volatility of the compounds, and to generate products with reduced polarity. Cholesterol derivatization is usually achieved by using trimethylsilyl compounds as reagents (silylation reaction). A common method for this purpose uses N,O-bis(trimethylsilyl-trifluoroacetamide/trimethylchlorosilane). Mass spectrometry (This section is derived from https://doi.org/10.1038/s41467-019-14180-4. Creative Commons Attribution 4.0 International License: http://creativecommons.org/licenses/by/4.0/). Mass spectrometry (MS) has become the method of choice for lipid analysis, offering label-free detection at high sensitivity and structural characterization capability. However, large-scale lipid analysis with a comprehensive capability of revealing all levels of structure information still represents a significant analytical challenge for lipidomics. General protocols, for instance, have five levels in terms of structure information, including lipid class, fatty acyl identities, fatty acyl sn-positions, and C=C location/geometry (viz cis/trans) in the fatty acyl. Successful attempts for determining C=C locations in fatty acyls or their sn-positions have already been reported for MS analysis, enabling the characterization of detailed structure moieties and identification of lipid structure isomers. An extremely useful feature offered by lipid isomer analysis is the relative quantitation achieved at high precisions without requiring the use of lipid standards, which are not readily available. Remarkably, our recent study demonstrated a close correlation between the lipid C=C location isomer compositions and Type II diabetes, which owes to tighter regulation on lipid desaturation, allowing efficient elimination of interferences due to variations among samples" Various methods have been explored for differentiating the lipid C=C location and sn-position isomers. Ozone-induced dissociation (OzID) and ultraviolet photodissociation (UVPD) have been used to determine both sn-positions and C=C locations in GPs. By coupling the Paternò-Bǜchi (PB) photochemical reaction with tandem MS (MS/MS), qualitative and quantitative analysis of lipids with C=C specificity from complex biological samples can be accomplished. PB reaction converts the C=C to an oxetane which can be preferentially fragmented by low-energy collision-induced dissociation (CID). An ideal analytical tool for lipidomics to survey a wide range of lipids in discovery work should not only provide detailed information at multiple lipid structure isomer levels (e.g., C=C location/geometry and sn-position), but also be feasible for large-scale quantitative analysis. UVPD is capable of assigning C=C locations and sn-positions of fatty acyls, while OzID may be the only one that has been well demonstrated for assigning C=C locations in sn-specific fatty acyls. One problem of OzID is the long reaction time required for the ion trap implementation, however recent work has demonstrated the ability to perform OzID on LC-compatible time scales in the high-pressure regions of the MS system. For the PB reaction method, both shotgun analysis and HPLC-PB-MS/MS workflow have been developed for identifying a large number of C=C location isomers. Figure \(5\) describes methods for the determination of C=C location and sn-isomers in lipids. 10.05: Problems 10.1 Questions 1) Explain why some lipid classes are membrane-forming and some are not. Give examples that support your logic. 2) a. Using the images of fatty acids below, write the symbolic name (N:N) for each. For the following use FA 1, 2, 3, or 4, some may have more than one answer. b. Which fatty acid melts at the highest temperature? c. Which fatty acid is a w-3 fatty acid and is essential for human growth and development? d. Which fatty acid, if found in a membrane bilayer, would increase membrane fluidity? e. Which fatty acid is the most unlikely to be found in nature, and why? 3) For the following questions, use the interactive micelle from Section 10.1: a. When looking at the micelle in “Ball and Stick” form, do you think the core (center of the micelle) could be accessible to water molecules? Explain. b. Examine the micelle under Style -> Surface Type -> Molecular Surface. Did your answer from part a change? Do there appear to be any solvent-accessible regions that water could use to enter the core? c. Finally, examine the micelle under Style -> Surface Type -> Solvent Accessible. Does this view change your answer to a.? Can you think of any medical or research applications in which a micelle could be useful given what you have found about its solvent accessibility? 4)Using the figure below, hypothesize what type of lipid could be represented by the geometrical shapes in the membrane. Why are the different shapes necessary for membrane formation? Could a membrane be formed with only one shaped lipid? If so, what, and explain why. 5) A cell membrane has the ability to remodel in response to stress in order to maintain membrane integrity. In the situations below, how could the membrane be remodeled to promote an intact structure? (I.e. what types of lipids could be added or removed to maintain a functional membrane? a. Increase in temperature b. Decrease in temperature 6) For each lipid below, name the type of lipid (membrane lipid, triacylglycerol, storage lipid, sphingolipid, wax, sterol, membrane glycerolipid, none of these), if it could be found in a membrane, and if it is a fatty acid or isoprenoid derived.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/10%3A_Lipids/10.04%3A_Working_with_Lipids.txt

Search Fundamentals of Biochemistry Introduction One easily understandable function of membrane bilayers is to separate the inside and outside of the cell or intracellular organelles. Yet as we mentioned before, such barriers can not be so rigid and impenetrable that they prevent the movement of materials across the membrane. Also, all cells must sense and respond to their environment through a process called signal transduction. We have already discussed lipid molecules involved in signaling. Now let's turn our attention to proteins that associate with the membrane and confer added functionalities to it. Figure \(1\) reviews some of the features of membranes we've discussed before and shows a simple bilayer (top) to the complicated membrane/cell wall of bacteria. Types of Membrane Proteins Although we presented this image earlier, Figure \(2\) reviews the details that should now be clearer to you. In this section, we will explore membrane proteins in more detail. Proteins can be loosely associated with the membrane (peripheral or extrinsic) or can embed deeply and most typically pass through the membrane and become a transmembrane (also called integral or intrinsic) protein. Sometimes they pass through using a single alpha helix, while other times they pass through multiple times (for example seven times in G-protein coupled receptors). They can also be classified based on the number of leaflets of the membrane they cross, as shown in Figure \(3\). Peripheral Proteins The proteins interact with a membrane through protein-lipid head group interactions, but might slightly penetrate the membrane. Those that do would be classified as monotopic peripheral. Peripheral proteins are generally easy to remove from a membrane in vitro by changing solution ions concentration as the interactions are often ion-ion nature. The first model below shows the binding of a matrix Metalloproteinase (MPP) 12 to a lipid bilayer. This protein is involved in inflammation, wound healing, arthritis, cardiovascular disease, and remodeling of neural synapses, suggesting a broad role in recovery from cell and tissue aberrations. MMP-12 is secreted by macrophages so it is considered a water-soluble (aqueous) protein. It travels to viral cells and appears to display activity not in aqueous solution but near membranes, implying activation of the enzyme through binding to the bilayer. Studies show the catalytic domain of MMP 12 can bind bilayers through both α- and β- secondary structure regions of the protein. Figure \(4\) shows an interactive iCn3D model of the protein and its interaction with the membrane through the alpha-helical region. Once bound to the membranes, catalytic activity increases. Other MMPs localize to membranes in other ways. MMP-7 interacts with heparan sulfate proteoglycans (CD44) and lipid rafts. Others bind transmembrane proteins like integrins. Other examples of peripheral protein include many precursor forms of protein clotting factors. Clotting is initiated when the serine protease thrombin cleaves fibrinogen to form fibrin, which self-associates to form a fibrin clot, or when thrombin activates receptors in platelets. The soluble precursor of thrombin, prothrombin, a zymogen, is activated on membrane binding through interactions with several proteins assembled on a negatively charged phospholipid (like phosphatidylserine) bilayer in the prothrombinase complex. How does the precursor zymogen interact with the membrane? It requires calcium ions, which bind to a series of gamma-carboxylated glutamic acid (GLA) residues on the zymogen. The enzyme that carboxylates the zymogen depends on Vitamin K. Figure \(5\) shows an interactive iCn3D model of bovine prothrombin Fragment 1 (N terminal) bound to a bilayer through its GLA domain (1NL2). The Gla sidechains are shown in CPK-colored sticks and interact with Ca2+ ions (gray spheres). Click on this link to see a zoomed view of just the calcium ions and Gla sidechains: https://structure.ncbi.nlm.nih.gov/i...xydtj8a4WHgdb7 The Gla domain in the absence of calcium ions is disordered. On binding, an ordered linear alignment of bound calcium ions is formed, stabilizing the ordered structure of the Gla domain and allowing interaction with the membrane. Three nonpolar amino acid side chains, Phe 5, Leu 6, and Val 9, are now clustered and exposed, allowing penetration of this hydrophobic patch part-way into the membrane. They are represented in cyan spacefill just underneath the surface of the red dots in the model above (the red dots are dummy atoms that represent the outer bilayer leaflet). Given this penetration, this protein domain would then be considered monotopic. What is not shown in the model is the role of negatively charged phosphatidyl serine. Studies have shown that the head group of serine in lysophosphatidylserine (which has only one acyl group) provides additional ion-ion interactions with the Ca2+ ions that also bind Gla residues 17 and 21. Arg 10 and Arg 16 also interact with the phosphatidyl serine head group. Phosphatidylcholine could also spatially fit into the active site but electrostatic interactions would prevent it. Why? Lipid-Anchored Proteins We have studied lipids, proteins, and carbohydrates. Although phospholipids can spontaneously form bilayers, the actual structure of biological membranes is made much more complicated through the addition of protein and carbohydrate substituents to the membrane. Soluble proteins can be made to insert into bilayers by the addition of nonpolar attachments. Localization to a membrane changes the functional expression of the protein. Several examples of such attachments are described below. Fatty acid linkers Two common covalent modifications of proteins are N-myristoylation (attached myristic acid - 14:0 - through an amide link) and S-palmitoylation (attached palmitic acid - 16:0 - through a thioester link with a Cys). Myristoylation is usually a cotranslational modification in eukaryotic and viral proteins that occurs after cleavage of the N-terminal methionine. Figure \(6\) shows an image of the serine/threonine phosphatase 2C (1A6Q) with its N-terminal glycine myristoylated. It should be obvious how this post-translationally modified protein interacts with a membrane. Figure \(6\): The serine/threonine phosphatase 2C (1A6Q) with its N-terminal glycine myrisoylated This modification is a key part of initiating immune system signal transduction pathways. The modification is catalyzed by N-myristoyltransferase (NMT) using myristoyl-coenzyme as the fatty acid acyl donor. This activates the function of the protein in part by reducing the dimensionality of substrate diffusion to the protein to the 2D surface of the membrane instead of a 3D search in the cytoplasm. NMT acylates protein at this consensus sequence: G1X2X3X4S/T5X6R7R8. Likewise, many signaling proteins are palmitoylated, leading to protein recruitment to membranes. Small G proteins like Ras, Rho, and the alpha subunit of heterotrimeric G proteins are often palmitoylated. This modification is also be found in transmembrane proteins in which localization is not an issue (see example xx below). In such circumstances, the modification might however help in targeting the proteins to rafts within the membrane. Palmitic acid is saturated and the addition of it to a protein might target it to more ordered regions of the membrane with cholesterol and sphingolipids within rafts. Isoprenoids linkers: The isoprenoids farnesyl (15C) or geranylgeranyl (20C)are added to a CAAX carboxy-terminal sequence in a target protein like RAS, where C is Cys, A is aliphatic, and X is any amino acid, which helps target proteins to the membrane. The enzymes used for these modifications are farnesyltransferase (FTase) and protein geranlygeranyltransferase I (GGTase I), respectively. For this and the other modifications, it has the potential to do more than target proteins to the membrane. The modification can also modulate protein-ligand interactions and protein stability. Ras, a key signaling protein, is a target of prenylation. Ras and other small G proteins are involved in a large percentage of human cancers. As the G protein Ras has somewhat of a billiard ball surface with obvious sites to target drugs that would affect its aberrant function in cancers, efforts have been made to target the prenyltransferases necessary to target it to the membrane. In humans, there are 3 different genes in the Ras family, H-Ras and N-Ras, whose gene products localize to both plasma and Golgi membranes, and K-Ras, which localizes predominantly to the plasma membrane. These and other G proteins bind GTP and possess GTPase activity. The GTP-bound form is active, while the GDP form is inactive. Point mutations that attenuate or prevent GTP cleavage leave the protein continually activated which contributes to oncogenesis. KRas has two predominant isoforms, 4A, the canonical form (also called 2A), and 4B (also called 2B) that arise from alternative splicing of the primary RNA transcript. The C-terminal protein sequences of isoform 4A and 4B differ significantly. Isoform 4A: QYRLKKISKEEKTPGCVKIKKCIIM Isoform 4B: KHKEKMSKDGKKKKKKSKTKCVIM The farnesylation motif site containing the modified Cys are highlighted in yellow above. The same cysteine is also often carboxymethylated. The Cys six residues from the farnesylated Cys in isoform 4A are also often palmitoylated Figure \(7\) shows isoform KRas 4B bound to a membrane bilayer through its farnesylated tail. (PDB file provided by Alemayehu (Alex) Gorfe. Viney Nair and Andrew McCammon). The tail is essential for its function at the plasma membrane where KRAS-mediated signaling events occur. Phosphodiesterase-δ (PDEδ) binds to KRAS4b and plays an important role in targeting it to cellular membranes. Note that the farnesyl attachments only penetrate part of the upper leaflet. Glycosyl-phosphatidylinositol linkers Normally soluble cytosolic proteins can become attached to membranes through the addition of a glycosyl phophatidylinositol (GPI). The attachment usually contains a conserved tetrasaccharide core of three mannoses (Man) and one unacetylated glucosamine (GlcN) linked to the carboxy terminus of the protein. The GPI can be further modified with extra galactoses and mannoses, as well as additions to the PI group, which secures the protein in the membrane. Figure \(8\) shows the common backbone for GPI anchors. Note the additions of the phosphoethanolamines to the core polysaccharide. GPIs are found in eukaryotic cells and link many surface antigens, adhesion molecules, and hydrolases to the membrane. GPIs from Plasmodium falciparum, the malarial parasite which kills about two million people each year, appear to act as a toxin and are the most common CHO modification of the parasite protein. Mice immunized against the GPI sequence, NH2-CH2-CH2-PO4-Man (α1-2) 6Man (α1-2) Man (α1-6) Man (α1-4) GlcNH2 (α1-6) myo-inositol-1,2-cyclic-phosphate, were substantially protected from malarial symptoms and death after they were exposed to the actual parasite. Figure \(9\) shows a cross-section of a membrane (with cholesterol, PE, SM) containing the glycosylated form of the human complement regulatory protein CD59 protein (1cdr) with a GPI anchor attached at its C-terminus. Note that the middle part of the anchor (glycan) holds the actual protein well above the top of the lipid bilayer. The soluble protein is also glycosylated. The protein binds to complement proteins C8 and/or C9, which are effector immune proteins that assemble on the surface of a cell undergoing lysis. The GPI anchor is shown in spacefill. Note that it only extends halfway into the bilayer, as you would expect from the size of the fatty acids attached to the phosphatidyl inositol. The glycan part of the GPI is shown in spacefill between the lipid and its protein attachment site. The protein is also glycosylated in the extracellular domain. Something new! A new (5/21) and totally unexpected type of glycosylated molecule has been found at the outer leaflet of mammalian cells - a glycosylated RNA, as shown in Figure \(10\). This adds RNA to the lipids and proteins as a target for glycosylation. These surface glycoRNAs interact with antibodies again ds-RNA and the Siglec lectin family. They are found in cells in vivo and cultured cells in vitro. Figure \(10\): A glycoRNA - a small noncoding RNAs with sialylated glycans. Park. https://doi.org/10.14348/molcells.2021.0178 www.molcells.org. Creative Commons Attribution-NonCommercial-ShareAlike 3.0 Unported License. To view a copy of this license, visit http://creativecommons.org/licenses/by-nc-sa/3.0/. Flynn et al., Small RNAs are modified with N-glycans and displayed on the surface of living cells, Cell (2021), https://doi.org/10.1016/j.cell.2021.04.023 Transmembrane (Integral) Proteins These proteins pass through the membrane either one in one pass, usually with a single alpha-helix or many membrane-spanning helices. For example, G protein-coupled receptors, often called serpentine receptors, cross the membrane seven times. There are three different types based on the number of types the protein crosses the membrane and the type of secondary structure used in crossing: biotopic (single pass), alpha-helical polytopic, and beta-barrel. These proteins are found in all types of membranes and have many types of functions, from receptors, receptor ligands, structural, adhesion, transport, gene regulation, and transport. Transmembrane Biotopic - Single Pass Proteins The are 4 types of single-pass transmembrane proteins: • Type I: N-terminal outside of the cell (extracellular) and the precursor signal sequence on the N-terminus which is a localization sequence is removed • Type II: N-terminal intracellular and with the transmembrane domain close to the N-terminus • Type III: N-terminus extracellular and no signal sequence in precursor protein • Type IV: N-terminus intracellular and the transmembrane domain close to the C-terminus The transmembrane domain of single-pass integral membrane proteins consists of a single alpha-helices with nonpolar side chains extending outward from the helical axis where they interact with the nonpolar lipid parts of the membrane. These nonpolar sides are more stable in nonpolar environments. To study such proteins in a less complex environment, membranes are often "dissolved" in nonpolar, single-chain amphiphilic detergents. These single-chain amphiphiles form micelles in the absence of membrane proteins but can form mixed micelles in which the nonpolar part of the protein is surrounded in the detergent micelle by the nonpolar acyl chains of the detergent. Figure \(11\) (top) below shows just the transmembrane and juxtamembrane (next to the membrane) domains of the single pass Notch protein, which is critical in many signal transduction pathways. The top images in the figure above show different ways to represent the protein in the bilayer, with the right-hand image showing a cross-section through the membrane to better show how the protein passes through the bilayer. The bottom images in the figure above show the protein after excess detergent, in this case, octylglucoside, is added to the protein-containing bilayer. Here are some examples of bitopic single-pass transmembrane proteins Cadherins All structures need support and connections. At the macro level, the skeleton supports the mass and organization of organs and tissues in whole organisms. Within an organ, how can cells hold together? How do they adhere to each other? Certainly not through outer leaflet lipid contacts as the outer surface of the leaflet is typically charged. The extracellular matrix does provide some of the glue that holds cells together. At a more detailed level, transmembrane proteins are involved. One class of adhesive proteins is cadherins, a calcium-dependent cell adhesion molecule. There are over 100 human cadherins. They are mostly ditopic, single transmembrane pass proteins. Their cytoplasmic domains interact with proteins like catenin, which then bind to the interior cytoskeletal network composed of actin and other proteins. This provides a way for the intracellular region to regulate the extracellular interactions of the cell. The extracellular domain is composed of five repeating "cadherin" domains, each around 110 amino acids, that can fold independently. Calcium ions bind at the domain interfaces. A cadherin can interact with other cadherin domains on other cadherins on other cells, leading to cell adhesion. Essentially, the receptor cadherin on one cell binds the ligand cadherin on the other. As metastatic tumor cells lose their adhering feature and leave the site of the primary tumor, you would expect that mutations in cadherins are often involved. They may also be involved in cell sorting during morphogenesis, "regulation of tight and gap junctions, and in the control of intercellular spacing". Figure \(12\) shows a "constructed" image of cadherin-1 (1L3W) interacting with cytoplasmic β-catenin (1I7X) through a modeled transmembrane helix (amino acids QIPAILGILGGILALLILILLLLLFLRR, amino acids 706-731). No full-length structure of cadherin in a membrane is available. Membrane Protein Kinases Kinases are enzymes that phosphorylate substrates. Hexokinase is a protein enzyme that catalyzes the phosphorylation of a hexose substrate such as glucose. A protein kinase is a protein enzyme that phosphorylates a protein substrate. That protein could be another copy of itself or another protein. We will see in Chapter 12 that many protein kinases are involved in cell signaling. Many tyrosine protein kinases are bitopic single-pass integral membrane proteins that become active on binding a ligand. Typically, on binding an extracellular ligand, two monomeric copies of the kinase form a dimer in the membrane, activating a tyrosine kinase cytoplasmic domain, which typically phosphorylates (using ATP as a substrate) the other member of the dimer in an "autophosphorylation" reaction. Sometimes the dimers are held together by disulfide bonds. Figure \(13\) shows a "constructed" image of the human dimeric insulin receptor. One of the monomers is shown in gray. The other monomer is shown in colors corresponding to the domain organization of the protein. Each extracellular dimer (6PXV) has two insulins bound (yellow spacefill). The intracellular domains (1IR3) are activated on insulin binding. No full-length structure of full insulin receptor in a membrane is available. Almost half of all helical membrane proteins in humans are bitopic, compared to between 20-25% in prokaryotes. Humans have 10-20 fold more bitopic proteins than E.Coli. There appear to be about 196 bitopic proteins in E. coli (located in the inner membrane ) and 70 in M. jannaschii (Archea in plasma membrane). In humans, 57% are in the plasma membrane, with the rest distributed between the Golgi, ER, nuclear, mitochondrial and chloroplast membranes. In single-celled yeast, only 8% are in the plasma membrane. Beta-Dystroglycan This protein is another example of a bitopic protein with a single alpha-helix membrane domain. Dystroglycan is a dimer of alpha and beta subunits. Alpha-dystroglycan is a peripheral protein that binds beta-dystroglycan, a transmembrane protein. Alpha dystroglycan also binds lassa virus and lymphocytic choriomeningitis virus glycoprotein, as it serves as viral receptors. It also binds the protein dystrophin, a protein missing in Duchenne muscular dystrophy, which affects 1 out of 5000 live male births. As an integral transmembrane protein, beta-dystroglycan connect the extracellular matrix to the cytoskeleton through dystrophin. Alpha- and beta-dystroglycan share the same gene, which codes one long protein which is proteolyzed post-translationally to form the alpha (N-terminal end) and beta subunits (C-terminal end). Figure \(14\) shows a schematic outline of dystrophin and the dystrophin-associated glycoprotein complex (DAGC). Figure \(14\): Schematic outline of dystrophin and the dystrophin-associated glycoprotein complex (DAGC). Dystrophin contains N-terminal (NT), middle rod, cysteine-rich (CR), and C-terminal (CT) domains. The middle rod domain is composed of 24 spectrin-like repeats (numerical numbers in the cartoon, positively charged repeats are marked in white color) and four hinges (H1, H2, H3, and H4). Dystrophin has two actin-binding domains located at NT and repeats 11-15, respectively. Repeats 1-3 interact with the negatively charged lipid bilayer. Repeats 16 and 17 form the neuronal nitric oxide synthase (nNOS)-binding domain. Dystrophin interacts with microtubules through repeats 20-23. Part of H4 and the CR domain binds to the β-subunit of dystroglycan (βDG). The CT domain of dystrophin interacts with syntrophin (Syn) and dystrobrevin (Dbr). Dystrophin links components of the cytoskeleton (actin and microtubule) to laminin in the extracellular matrix. Sarcoglycans and sarcospan do not interact with dystrophin directly but they strengthen the entire DAGC, which consists of dystrophin, DG, sarcoglycans, sarcospan, Syn, Dbr, and nNOS. .Disease Models & Mechanisms (2015) doi:10.1242/dmm.018424vailable via license: CC BY 3.0 Transmembrane - Alpha-helical polytopic There are so many intriguing examples of these proteins. We'll illustrate just two. Rhodopsin-like receptors and pumps These proteins are involved in cell signaling and are the target of most pharmaceutic drugs. G protein-coupled receptors (GPCRs) are incredibly important and we will discuss them extensively in Chapter 12. GPCRs are cell receptors that span the membrane seven times in a serpentine fashion. They bind ligands (neurotransmitters, hormones, etc) in the extracellular or internal membrane domains (the latter for hydrophobic ligands), and through propagated conformations changes alter the cytoplasmic domain where they functionally interact with a heterotrimeric G protein. Figure \(15\) shows an interactive iCn3D model of the human cannabinoid receptor with bound cholesterol and Δ 9 -tetrahydrocannabinol (Δ9 -THC) in spacefill (5xra). The red dummy atoms represent the outer leaflet and the blue the inner. Δ 9 -THC is a partial agonist and tunes the response of the receptor. The active site is conformationally somewhat flexible or plastic. Other ligands bound to it act as antagonists instead of agonists and must do so by eliciting nonactive conformations. ABC Transporter Figure \(16\) shows an interactive iCn3D model of the P-glycoprotein multidrug resistance transporter protein (6nf1). The spacefill ligands represent Zosuquidar, which binds with high affinity to P-glycoprotein and inhibits its activity, making it a cancer agent as it prevents chemotherapeutic drugs that have entered the cell from being pumped out. The protein chain interacting with it on the cytoplasmic face is an antibody fragment used to stabilize the P-glycoprotein so crystals could form. Transmembrane Beta-barrel transmembrane We will focus on two of these proteins. Outer Membrane Factor (OMF) - Gram-negative bacteria Figure \(17\) shows an interactive iCn3D model of a beta-barrel transmembrane protein OPRM - Outer Membrane Factor (4y1k) from Pseudomonas aeruginosa that acts as a pore. It also has a palmitoyl fatty acid in thioester linkage to Cys 1 of the protein for extra but unneeded anchorage. Use this link for another view: https://structure.ncbi.nlm.nih.gov/i...FtY5FL7&t=4Y1K (OPM) in iCn3D This protein is part of a large complex of proteins that spans both the inner and outer membranes of Gram-negative (examples E. Coli and Pseudomonas aeruginosa) bacterial cell walls. Unfortunately for humans, this protein complex pumps out toxins (to the bacteria) like antibiotics, which makes bacteria resistant to these drugs. The OPRM acts as the outer passageway or duct for the pumped molecules. The Bacterial Outer Membrane Factor (OMF) protein differs in sequence but all form the beta-barrel duct. The E. Coli version of OMF has a triacylated lipid modification of the N-terminus. The N terminal lipid modification might be necessary for the initial attachment of the protein to a membrane before the insertion of the beta-barrel. As such, the enzymes involved in the attachment of the tail could be targets for new antibiotics. Voltage-dependent anion channel (VDAC) - mouse This protein regulates the movement of molecules between the cytoplasm and the interior of the mitochondria across the outer mitochondrial membrane. VDAC also serves as a docking site or scaffold for the assembly of molecules into a complex that regulates mitochondrial function. The protein's conformation and hence function are regulated by changes in the transmembrane potential, which we will explore in the next sections. Hence the protein and its function are voltage-dependent. Figure \(18\) shows an interactive iCn3D model of mouse VDAC with a beta-barrel formed by 19 beta-strands (3emn). Note the N-terminal alpha-helix resident inside the channel opening. This helical section moves on changes in membrane potential, gating open and hence regulating the flow of metabolites and ions across the membrane through the pore. At a low transmembrane potential (10 mV), the conductance is high as the channel is in the open state. When the potential increases to 30 mV (either + or -) conductance drops as the protein forms the closed state. Now that you understand the structure of membrane proteins, let's explore a key type of function of a subset of integral membrane proteins: the movement of molecules/ions across the membrane.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/11%3A_Biological_Membranes_and_Transport/11.01%3A_Membrane_and_Membrane_Proteins.txt

Learning Objectives • Understand the structure and function of biological membranes. • Describe the different types of diffusion and how they operate across a membrane. • Explain the concept of concentration gradient and how it drives diffusion. • Describe the mechanisms of passive and facilitated diffusion, including their differences and similarities. • Understand the role of carrier proteins and channels in facilitated diffusion. • Describe the factors that can affect the rate of diffusion across a membrane. • Understand the importance of membrane transport in maintaining homeostasis in cells and organisms. Search Fundamentals of Biochemistry Diffusion Across a Membrane We have studied molecular aggregates (micelles and bilayers) and macromolecular structures (mostly proteins). We also studied binding interactions which are the first steps in the expression of the biological activity of a macromolecule. For some proteins, reversible binding is their sole function (consider the binding of dioxygen to myoglobin and hemoglobin). For many others, it is not. For those, what can happen next? You already have one possible answer. A bound reactant, which we will call a substrate, can be converted to a product in a chemical step involving the breaking and making of covalent bonds catalyzed by a protein enzyme. However, there is an even simpler process that does not involve covalent bond changes. If a small molecule is bound to a membrane protein, it could be transported in a purely physical step across a membrane. Just as reactions can proceed with and without an enzyme, a solute can move down a concentration gradient across a semi-permeable membrane, driven by diffusion alone in a thermodynamically favorable process, either by itself, in a process called passive diffusion, or with the assistance of a membrane protein, in a process called facilitated diffusion. Large pores made of assemblies of proteins can also be formed that allow the passage of many solutes across the membrane. There are many occasions when it would be optimal to move a molecule across a membrane from a region of low to high concentration. This process is called active transport. It is not thermodynamically favored so it requires an external energy source. This is often the thermodynamically favored cleavage of ATP to ADP and Pi. The uphill transport can also be powered by the downhill diffusion of a "co-transport" molecule from high to low concentration. We will explore all of these processes in this and the remaining chapter sections. First, we should understand the simplest process, "passive diffusion", that requires no protein "help". Let's step back and think about how difficult it is for a chemical species to cross a lipid bilayer. Chemical intuition would tell us that both size and polarity are important. The bigger the size and the greater the charge, the more difficult it would be to cross the membrane. The permeability coefficient is related to the ease with which solutes traverse the membrane. Figure $1$ shows the permeability coefficients for relevant biological molecules. Smaller, higher charge density ions (like Na+) have a lower permeability coefficient than do larger, lower charge density ions (like K+) as seen in (Table $1$). What about natural membranes? Table $1$: Permeability coefficient (cm/s) of natural and synthetic membranes to D-Glucose and D-mannitol at 25 °C Membrane Preparation D-Glucose D-Mannitol Synthetic Lipid Bilayer 2.4 x 10-10 4.4 x 10-11 Calculated Passive Diffusion 4 x 10-9 3 x 10-9 Intact Human Erythrocyte (red blood cell) 2.0 x 10-4 5 x 10-9 Looks like D-glucose gets a little help in getting across. We'll see the mechanism below. Passive Diffusion Let's start with the passive diffusion of uncharged solute A across a membrane, which can be represented by the chemical equation Aout ↔ Ain. Intuitively, you probably believe that the rate of net diffusion or the flux of A across the membrane is directly proportional to the concentration gradient across the membrane. If concentrations of A are identical across the membrane, the net flux J should be 0. If you double the concentration gradient, the net rate should double. We will see that the net rate is a linear function of [ΔA] across the membrane. Figure $2$ shows the flux of Aout across a semipermeable membrane of thickness Δx (we will use dx instead when Δx is very small). Let's animate diffusion using a PHET simulation, as shown in Figure $3$. Figure $3$: PHET animation of passive diffusion. PHET: https://phet.colorado.edu/ The flux of molecules $A$ ($J_A$) is proportional to the concentration gradient across the membrane, $ΔA/Δx$ (which we will refer to as $dA/dx$ which is the derivative of $A$ with respect to $x$). The equation below is Fick's First Law of Diffusion: $\mathrm{J}_{\mathrm{A}} \propto \frac{d \mathrm{~A}}{d x}=-\mathrm{D} \frac{d \mathrm{~A}}{d x} \label{Ficks1}$ where $D$ is the diffusion coefficient. The negative sign is necessary since concentration increases to the left in the figure above in the opposite direction of net flux which is to the right. For these derivations, we will assume the JA is the initial flux. That is, the flux is measured for a short enough time that the relative concentrations of A on both sides of the membrane do not change significantly. It should be clear that eventually the net flux levels off to zero when the concentrations of A on both sides of the membrane are equal. Under these conditions, the free energy $G_{A\, out}$ = $G_{A\, in}$, so $ΔG = 0$. This thermodynamic relation can also be expressed as J_A=-L \frac{d G_A}{d x} This equation bridges the kinetic and thermodynamic aspects of diffusion. Dimensional analysis of Fick's 1st Law (Equation \ref{Ficks1}) shows that the units of $D$ are cm2/s which gives the number (dimensionless) of molecules crossing a 1 cm2 surface area of membrane each second. $J$ = moles/area/sec = mol/cm2.s = - (cm2/s) mol/cm3/cm. Hence the units of $D$ are cm2/s. Let's rearrange Fick's 1st Law and use a bit of calculus to get Equation 4. \begin{gathered} \mathrm{J}_{\mathrm{A}} \int_0^{\mathrm{x}} \mathrm{dx}=-D \int_{\mathrm{A}_{\text {out }}}^{A_{\mathrm{in}}} \mathrm{dA} \ \mathrm{J}_{\mathrm{A}} \mathrm{x}=-\mathrm{D}\left(\mathrm{A}_{\mathrm{in}}-\mathrm{A}_{\mathrm{out}}\right) \ \mathrm{J}_{\mathrm{A}}=\frac{\mathrm{D}}{\mathrm{x}}\left(\mathrm{A}_{\text {out }}-\mathrm{A}_{\mathrm{in}}\right) \end{gathered} or J_A=P\left(A_{\text {out }}-A_{\mathrm{in}}\right)=P \Delta A where P is the permeability coefficient, which has units of cm2/s/cm or cm/s. (We discussed permeability coefficients for different solutes traversing model bilayers when we discussed lipids.) That unit is less intuitive to understand but the final unit is very intuitive. A plot of JA vs (Aout - Ain) is linear, with a slope of P = D/x It's important to remember that there is still diffusion of A across the membrane at equilibrium since the equilibrium is dynamic. There is no net diffusion, however. This is where animations come in handy. The table below shows the reaction diagram (left), graphical results of the progress curve(middle), and animations for the reversible diffusion Aout ↔ Ain across a membrane with the conditions shown below. The reactant A and product are called species and are shown as green spheres. The yellow square is a reaction node indicating a reaction connects A to P. The lines connect the species that participate in the reaction. The velocities (slope of the concentration vs time curve at any given time) are called fluxes, J, in Vcell and many other similar programs. When we get to metabolism, we will talk about fluxes of metabolites through pathways. Also, fluxes are used to describe the rate of movement of solute through membranes. VcellModel Initial Conditions Click Select Omex below to run the simulation produced in Vcell. Animations Now let's look at animations of the same passive diffusion reaction of a neutral species across a semi-permeable membrane. Animations are by Shraddha Nakak and Hui Liu. Aout ↔ Ain [Ain]t=0 = 100; [Aout]t=0 = 0; P = 2 Aout ↔ Ain [Ain]t=0 = 50; [Aout]t=0 = 0; P = 2 Note the dynamic nature of the diffusion. An equilibrium is reached when the number of particles inside equals those outside. Diffusion occurs in both directions from compartments of equal volume so that the particles moving to the outside don't escape into a comparatively huge volume. (Animations by Shraddha Nayak and Hui Lui) Passive Diffusion of ions across a membrane - Transmembrane Potentials Before we move on to facilitated diffusion, let's alter the scenario a bit and use a charged solute. In the example of passive diffusion above, the only thermodynamic driving force for the movement of A across the membrane was the ΔGA, the change in free energy/mol of A across the membrane (or more strictly Δμ = change in chemical potential). Solute A moves spontaneously across the membrane from high to low concentration. But what if A was charged? We could add a bunch of positively or negatively charged species to Figure $2$ and ask what would happen. You can't go to a chemical stockroom and find a bottle of K+ ions but you could find a bottle of neutral KCl. Let's make our experiment system a vesicle that has 0.1 M KCl in the aqueous inner compartment with 0.1 M NaCl on the outside. We could easily prepare such vesicles by making large unilamellar vesicles (LUVs) with entrapped 0.1M KCl in a solution of 0.1 M KCl and then separate the vesicles from the 0.1 M KCl not encapsulated on liposome formation using a size exclusion gel chromatography column equilibrated and eluted in 0.1 M NaCl. These vesicles are illustrated on the left of Figure $4$. In these prepared vesicles is there a net thermodynamic driving for K+ and Cl- to move from inside to outside the vesicle? Not for Cl- since its concentration is the same on both sides of the membrane (see Fick's 1st Law above). However, there is a clear thermodynamic driving force to take K+in → K+out. If the membrane was impermeable, net outward flux would not occur even though it is favored thermodynamically. Think of this as an example of a reaction under complete kinetic control! Note that in this example there is also a net driving force to move Na+ ion from outside to inside as well. In our next step, let's make the membrane permeable to only K+ ions. We can do this by adding a small antibiotic, valinomycin, which binds in the membrane and once there binds and carries K+ ions across the membrane. It is called an ionophore. Figure $5$ shows an interactive iCn3D model of K+ bound to Valinomycin. Valinomycin, from Streptomyces fulvissimus, is a cyclic peptide consisting of L and D-Val along with L-lactate and D-hydroxyisovalerate, connected through both ester and amide bonds​. The K+ ion is in the center. The six valine carbonyl oxygens bind the K+ ion. The hydrophilic groups are pointed toward the center, while the hydrophobic groups point to the outside of the structure, allowing the K+ ion to be sequestered in a polar environment as the nonpolar exterior of the complex passes through the membrane. This ionophore is specific for K+ and binds the smaller Na+ ion weakly. This can be accounted for by two factors. The smaller sodium ion doesn't bind as tightly to the chelating carbonyl oxygens. Also, the sodium ion has a higher charge density, so the Na+/water interactions must be more stable and more difficult to break than those to K+. The ion must be desolvated before it binds to the complex. Other ionophores are specific for other ions. Once the ionophore is bound, the kinetic barriers to K+ efflux are removed and it starts moving K+ from inside to outside. However, as soon as it does, the charge balance across the membrane is lost, with the outside becoming net positive and the inside becoming net negative. A transmembrane electric potential develops across the membrane. This disfavors stops K+ efflux to the outside and eventually stops it even as the concentration difference of K+ across the membrane still favors efflux. If you were a positive ion stuck in the middle of a membrane, as illustrated in Figure $6$, which way would you move? There are now two thermodynamic driving forces for K+ movement from inside to outside: • a ΔGconcentration which favors K+ efflux. At time t=0, ΔGconcentration << 0 and it becomes a bit less negative (less favored) with efflux. • a ΔGmembrane pot which is zero to start and slowly becomes positive, increasingly disfavoring K+ efflux. When these driving forces are equal and opposite, net K+ movement across the membrane stops and the system is in dynamic equilbrium. Since we use electric potential to describe electrical phenomena (electron, ion movement), we often use the word chemical potential in this case to describe the movement of ions across a concentration gradient. Add them together and we call it the electrochemical potential. \begin{array}{c} \Delta \mathrm{G}_{\text {electrochemical }}=\Delta \mathrm{G}_{\text {chemical }}+\Delta \mathrm{G}_{\text {electrical }} \text { or } \ \Delta \mu_{\text {electrochemical }}=\Delta \mu_{\text {chemical }}+\Delta \mu_{\text {electrical }} \end{array} We can use this understanding to derive an equation for flux J of a charged solute across a membrane of a given potential. The equation is called the Goldman Equation and is shown below. J=\frac{P \frac{Z F}{R T}\left(E_{1}-E_{2}\right) C_{1}\left(1-\frac{C_{2}}{C_{1}} e^{\frac{Z F}{R T}\left(E_{2}-E_{1}\right)}\right)}{1-e^{\frac{Z F}{R T}\left(E_{2}-E_{1}\right)}} where • P is the permeability coefficient • Z is the charge or valence on the ion • F is the Faraday constant • R is the ideal gas constant • T is temperature • E2-E1 and the reverse is the transmembrane potential • C2-C1 are the concentrations of the ions across the membrane Compare this to the Nernst equation which you learned in introductory chemistry courses. E=E^{o}-\frac{R T}{n F} \ln Q that relates the reduction potential of an electrochemical reaction to the standard electrode potential, temperature, and concentration where E is the potential difference. Now let's run a Vcell simulation for the diffusion of an anion across a semipermeable membrane. To do so we must first set the initial transmembrane potential to solve the Goldman equation numerically. At present, this type of simulation can not be embedded into this book. So instead, the concentration vs time graphs for two different simulations, one at an initial transmembrane potential (E2-E1) = - 0.001 (i.e. 0) and one at -60 mV (a typical cell resting potential), are presented in the Figure below. In each, the reaction is A-in↔ A-out [A-in]t=0 = 100. A-in↔ A-out [A-in]t=0 = 100; [A-in]t=0 = 0; P = 100; Vinitial = 0 (-0.001 V) A-in↔ A-out [A-in]t=0 = 100; [A-in]t=0 = 0; P = 100; Vinitial = -60 mV Here are animations that show the selective reversible movent of anions (A-, left panel, red) and cations (C+, right panel, cyan) as they move across a membrane from the inside to the outside. This is a very simplified simulation as it shows no counter ions on either side. Assume they exist as it would be impossible to have a "container" with just anions or cations. The initial transmembrane potential (t=0) is 0 (actually -0.001 to allow the calculations using Vcell). A-in ↔ A-out [A-in]t=0 = 100; [A-out]t=0 = 0; P = 100; Vinitial = 0 (-0.001 V) C+in ↔ C+out [C+in]t=0 = 100; [C+out]t=0 = 0; P = 100; Vinitial = 0 (-0.001 V) As anions move to the outside (left animation), the inside becomes less negative with respect to the outside, so the membrane potential V becomes more positive. This is indicated by the membrane changing to a blue color. Conversely, as cations move to the outside (right animation), the inside would become more negative with respect to the outside, so the membrane potential V becomes more negative. This is indicated by the membrane changing to a red color. (Animations by Shraddha Nayak and Hui Lui) Facilitated Diffusion Now let's return to the diffusion of a noncharged solute down a concentration gradient (i.e. favored) after binding to a membrane receptor. The answer to that question depends on the biological function of the macromolecule. We can simplify this process by adding one additional step as reflected in the equilibrium binding expression shown below: $\ce{M + L <=> ML <=> M + X} \nonumber$ This expression indicates that the free ligand has changed in some fashion to x. In the next two chapters, we will consider two kinds of transformations: • L is a ligand on the outside of a biological membrane (Lout) that binds to a membrane protein receptor, R. This undergoes a conformational change (as we studied in the binding of dioxygen to hemoglobin) which leads to the expulsion of the bound ligand to the inside of the membrane (Lin). This can be modeled with the simple equation: $\ce{R + L_{out} <=> RL <=> R + L_{in}}. \nonumber$ This process is called facilitated diffusion and represents a physical as opposed to chemical process since no covalent bonds are made or broken. This process proceeds down a concentration or chemical potential gradient (Δμ < 0) and hence is spontaneous (thermodynamically favored). If the ligand concentration is higher inside the cell, net diffusion moves it to the outside of the cell. Passive (non-facilitated) diffusion is kinetically slow in the absence of a receptor since membranes present formidable barriers to the passage of polar molecules. • L is a ligand (or substrate S) that binds to a protein enzyme, E. The bound substrate is chemically altered to produce a new product, P, which dissociates from the enzyme. This can be expressed most simply as: E + S <==> ES <==> E + P . Consider the mechanism illustrated in Figure $7$. Let's assume that for this system the initial flux will be measured. We would like to derive equations that show J as a function of Aout (assuming that Ain is negligible over the time course of measuring the initial flux. Also, assume that the J facilitated is much greater than J passive. In contrast to passive diffusion, JA is not proportional to Aout but rather to [Abound]. Consider this example to help you understand that proportionality. Pretend that the receptor is a truck that can carry one particle across the membrane at a time (i.e. 1/1 stoichiometry). Also, assume that the particle can't get across without being carried by the truck. If there are no trucks in the membrane, no load can be delivered. If there are trucks in the membrane but no particles in them, no load will be delivered. As the number of particles available to be loaded into the truck increase, the truck will have an increased chance to be loaded (depending of course on the affinity of the particle for the truck). If the number of loaded trucks is doubled, the number of particles dumped to the other side will double. Therefore, by analogy, JA is proportional to [RA], or J_A=\operatorname{const}[\mathrm{RA}]=\mathrm{k}_3[\mathrm{RA}] How can we calculate RA when we know A and R? Let us assume that Atotal (A0) is much greater than R0, as is the likely biological case, and Ain = 0. We can calculate RA using the following equations, and the same procedure we used for the derivation of the binding equation [\mathrm{ML}]=\frac{\left[\mathrm{M}_0\right][\mathrm{L}]}{\mathrm{K}_{\mathrm{D}}+[\mathrm{L}]} The equation for the dissociation constant KD \mathrm{K}_{\mathrm{D}}=\frac{[\mathrm{A}]_{\mathrm{eq}}[\mathrm{R}]_{\mathrm{eq}}}{[\mathrm{RA}]_{\mathrm{eq}}}=\frac{(\mathrm{A})(\mathrm{R})}{\mathrm{RA}} The equation of mass balance of R \mathrm{R}_0=\mathrm{R}+\mathrm{RA} \text { so } \mathrm{R}=\mathrm{R}_0-\mathrm{RA} Since we will assume that A0 is much greater than R0, we will not need the mass balance for A (which is Ao = A + RA). Substitute x into x and rearrange to get: \begin{gathered} \mathrm{K}_{\mathrm{D}}(R A)=(A)(\mathrm{R})=(A)\left(\left(\mathrm{R}_0\right)-(A)(R A)\right. \ \mathrm{K}_{\mathrm{D}}(R A)+(A)(R A)=(A)\left(\mathrm{R}_0\right) \ \left(\mathrm{K}_{\mathrm{D}}+\mathrm{A}\right)(R A)=(A)\left(\left(\mathrm{R}_0\right)\right. \ (R A)=\frac{\left(\mathrm{R}_0\right) A}{\mathrm{~K}_{\mathrm{D}}+\mathrm{A}} \end{gathered} Substitute x into z gives the final equation, \mathrm{J}_{\mathrm{A}}=\mathrm{k}_3[\mathrm{RA}]=\frac{\mathrm{k}_3\left(\mathrm{R}_0\right) A}{\mathrm{~K}_{\mathrm{D}}+\mathrm{A}}=\frac{\mathrm{J}_{\max } A}{\mathrm{~K}_{\mathrm{D}}+\mathrm{A}} It should be clear to you from this equation that: • a plot of JA vs A is hyperbolic • JA = 0 when A = 0. • JA = Jmax when A is much greater than KD • A = KD when JA = Jmax/2. These are the same conditions we detailed for our understanding of the binding equation. This derivation is based on the assumption that the relative concentrations of A, R, and AR can be determined by the KD for the interactions and the concentrations of each species during the early part of diffusion (i.e. under initial rate conditions). Remember under these conditions, Aout does not change much with time. Is this a valid assumption? Examine the mechanism shown in the above figure. Aout binds to R with a second order rate constant k1. RA has two fates. It can dissociate with a first-order rate constant k2 to Aout + R (to give the original species), or dissociate with a first-order rate constant of k3 to give Ain + R (as A moves across the membrane). If we assume that k2 >> k3 (i.e. that the complex falls apart much more quickly than A is carried in), then the relative ratios of A, R, and RA can be described by KD. Alternatively, you can think about it this way. If A binds to R, most of A will dissociate, and a small amount will be carried across the membrane. If this happened, then R is now free, and will quickly bind Aout and reequilibrate. This occurs since the most likely fate of bound A is to dissociate, not to be carried across the membrane, since k3 << k2. Vcell Model: Initial Condition Click Select Omex below to run the simulation produced in Vcell. Note that the JM values for facilitated diffusion are 1000 times the k values for passive diffusion "Receptors" in Facilitated Diffusion Two types of proteins are involved in facilitated diffusion, carriers and channels. Carrier proteins (also called permeases or transporters) such as the glucose transporter (GLUT1) move solute molecules across a membrane while channels/pores facilitate the diffusion of ions down a concentration gradient by providing a pore in the membrane. We won't describe in this section the more complicated processes of phagocytosis and endocytosis. These processes are illustrated in Figure $8$. In the case of permeases and transport proteins, ligands bind and induce a conformational change in the receptor as illustrated in the case of the glucose transport protein shown in Figure $9$. In channels and pores, a ligand can bind to the receptor (channel protein), which induces a conformational change in the receptor, a "ligand-gated" channel through the membrane. This process would lead to the diffusion of many ions across the membrane (down a concentration gradient) until the channel closes (which can be induced by ligand dissociation or other events). The mathematics we derived for the carrier proteins does not apply to the channel proteins. In addition, there are other ways to "gate" open a channel protein, which we will discuss later. Also, some transporters can move solute molecules across a membrane against a concentration gradient. These proteins require an external energy source (like ATP or coupling to the favorable collapse of a second transmembrane gradient) to drive this thermodynamically unfavored process. This is called active transport and will be discussed in the next chapter section. Both links above are from the Theoretical and Computational Biophysics group at the Beckman Institute, University of Illinois at Urbana-Champaign. These molecular dynamic simulations were made with VMD/NAMD/BioCoRE/JMV/other software support developed by the Group with NIH support. Carrier proteins (permeases or transporters) Now let's look at some examples of carrier proteins: Glucose Transport Proteins Glucose is a key metabolic fuel so its movement into cells is critical and hence highly regulated. There are multiple types of glucose transporters. GLUT 1, a plasma membrane protein, found in most cells, is responsible for constitutive or basal glucose uptake while GLUT 4 is involved in insulin-regulated uptake in skeletal and heart muscles and adipose cells, of glucose after meals. Its official name is solute carrier family 2 or, facilitated glucose transporter member 4. No structure is yet available for GLUT 4 but there is for GLUT1, which is highly expressed in cancer cells that have high energy demands. Figure $10$ shows an interactive iCn3D model of a glucose transporter, GLUT1 (5eqg), bound to an inhibitor, cytochalasin B (spacefill). The inhibitor binds in the inward-open state where glucose binds. Mitochondrial ADP-ATP Carrier Protein We will see in a few chapters that most of the ATP made in cells takes place in the mitochondrial matrix. It won't do cells much good if it stays in there since it is needed in the cytoplasm and elsewhere to drive unfavored processes. Likewise, when ATP is depleted in a cell, ADP is concomitantly high. What is needed is an inner mitochondrial membrane protein that can shuttle ATP out of the mitochondria and ADP in down concentration gradients. It would not make sense to need to power an uphill movement of ATP into the mitochondria from low to high concentration driven by ATP cleavage. Let's look at the structure of the bovine ADP-ATP carrier protein which resides in the inner membrane. Figure $11$ shows an interactive iCn3D model of the bovine mitochondrial ADP-ATP carrier protein (1okc). The transmembrane domain contains six alpha-helices which form a depression leading to the inner leaflet. The cyan spacefill amino acids on the bottom of the depression are RRRMM, which is a motif found in nucleotide carrier protein. A conformational transition must transiently open the depression into a channel. The spacefill molecule in CPK colors represents carboxyatractyloside, a diterpene glycoside that inhibits the carrier protein. A Special Case: Fatty acid carrier proteins You might guess that free fatty acids, derived for example from lipids after the actions of lipases on triacylglycerol, would not need a carrier protein to move across the cell membrane since they are almost completely nonpolar. Hexanoic acid can indeed pass readily, but for solute diffusion across the membrane, size matters as well. A whole family of proteins, Fatty Acid Transport Proteins (FATPs) have evolved to help long-chain fatty acids across membranes. Human fatty acid transport proteins are transmembrane proteins. Its mechanism of action is unclear. No crystal structures of these are readily available. Many proteins in this class catalyze the formation of fatty acid-CoASH derivatives, which is an endergonic reaction powered by ATP. The mechanism of fatty acid movement across the membrane probably may involve simple diffusion coupled to processes driven by ATP. However, there is still controversy on the role of passive vs facilitated diffusion for fatty acids. First, let's consider the problems facing a cell in moving fatty acids across two aqueous environments. Figure $12$ shows a mass balance depiction of the reservoirs of fatty acid in the extracellular and intracellular environment. Free fatty acids are very insoluble in aqueous solution so their concentrations on either side of the membrane are very low, in the low nanomolar range. Hence there is no great thermodynamic drive to move free fatty acids across the membrane. If you assume that fatty acids can reasonably transverse the membrane without a carrier protein, there would be no huge kinetic barriers to movement except their low concentrations. On each side of the membrane, the free fatty acids are in an "equilibrium" with protein-bound amino acids. In the blood and interstitial fluids, albumin, which can bind multiple fatty acids, is in high concentration, so it can act as a buffer to keep free fatty acids in a useful concentration range. Likewise, in the cytoplasm, fatty acid binding proteins (FABPs), which typically bind just one fatty acid, are also relatively high in concentration and buffer the free fatty acids in the cytoplasm. One other note. Free fatty acids are single-chain amphiphiles which makes them detergents, which could easily lyse cell membranes, so their free concentrations must be kept very low compared to the critical micelle concentration. Figure $13$ shows an interactive iCn3D model of the human brain fatty acid-binding protein bound to docosahexaenoic acid (1fdq). How do long-chain fatty acids cross the membrane? We will first examine the role of proteins. Let's look at a couple of players. Fatty acid transport proteins (FATPs): There are six members of the human FATP family, which is also known as the Solute carrier family 27. FATP 1 (SLC27A-1) is found in plasma and endoplasmic reticulum membranes and based on sequence analysis it is a single-pass membrane protein. Its highest expression is in muscle and adipose cells. It possesses a C-terminal AMP binding domain and acyl-CoA synthase activity. The mouse protein has an N terminal transmembrane domain and predictions from the human sequence show there is likely just one transmembrane helix at the N terminus (amino acid 13-35). Given that, and the absence of a 3D structure, it would appear that this protein would not bind and transfer the bound lipid across the membrane through a conformational change in the transmembrane domain. FATP 4, located in the ER membrane, is predicted to have 2 transmembrane helices, which still would probably be inadequate to serve as a class translocase. It is expressed in the endoplasmic reticulum cell membrane. Platelet Glycoprotein 4 (CD36): Another candidate is platelet glycoprotein 4, which is also called CD36, Glycoprotein IIIb, fatty acid translocase, or the thrombospondin receptor. The protein has many functions and binds many types of proteins (thrombospondin, fibronectin, collagen or amyloid-beta) and lipids (oxidized low-density lipoprotein (ox-LDL), anionic phospholipids, long-chain fatty acids, and bacterial diacylated lipopeptides). It is present in plasma and Golgi membranes. Sequence analysis shows that it passes through the membrane twice (amino acids 7-29 and 441-463) and it also is palmitoylated at both N- and C-terminal ends. A PDB structure (5LGD) for most of the protein except the putative N- and C-terminal helices are known. In the structure, it is bound to a malarial protein (shown in grey) and two palmitic acids (spacefill) bound in the nonmembrane domains. Again, from this description, it doesn't appear that the bound fatty acids are translocated via a conformational change in the receptor as described above for glucose and ADP/ATP. Fatty Acid Binding Proteins (FABPs): These proteins might also take part in the process. In addition to the cytoplasmic form, there is also a plasma membrane-associated fatty acid-binding protein (FABPpm), also known as FABP-1. It's the same protein as mitochondrial aspartate aminotransferase (UniProtKB - P00505 (AATM_HUMAN). It has many possible functions. Figure $14$ shows a possible model for how fatty acids may transfer or be handed off from albumin to membrane-bound GP36 or FATP-1, possibly through an intermediary protein like FABPpm (for GP36). Some proteins (albumin, FABP) deliver fatty acids to the membrane proteins (CD36, FATP-1), which deliver free fatty acids into the outer leaflet, where they flip to the inner leaflet, where they are picked up by membrane-associated cytoplasmic fatty acid binding proteins (FABP). Until structures are known for the transmembrane-bound proteins GP36 and FATP-1, whose full amino acid sequences don't suggest a classic carrier protein, this mechanism is a reasonable one. FATP-1 has acyl-CoA synthase activity so it is likely that the transferred fatty acid is converted to the acyl-CoASH before movement to the cytoplasm. Fatty acid transporters are also implicated in insulin resistance and type 2 diabetes. The alternative model proposes that long-chain fatty acids, the preferential energy source for cardiac muscle, can cross the membrane by passive diffusion (red boxed area above) even if the activity of CD36 is inhibited. Diffusion depends on an alteration of the pKa of outer membrane adsorbed free fatty acids (around 7.5) compared to free fatty acids in solution (around 4.5). The protonated fatty acid adsorbed to the membrane would move into the outer leaflet where it would flip to the inner leaflet and be picked up by cytoplasmic and/or peripheral membrane proteins. This process would be associated with movement of H+ across the leaflet as well. This type of diffusion has been observed in protein-free lipid vesicles and cells. Likewise, long chain fatty amines (instead of carboxylic acids) can diffuse into vesicles and cells which would support this passive diffusion if fatty acid-binding proteins don't bind the amine forms.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/11%3A_Biological_Membranes_and_Transport/11.02%3A_Diffusion_Across_a_Membrane_-_Passive_and_Facilitated_Diffusion.txt

Search Fundamentals of Biochemistry Introduction If you punched a hole or pore in the membrane, depending on its size, multiple types of chemical species could flow through it simultaneously. We'll talk about pores in the next section. Let's focus on channels, which have much smaller openings, which are gated open to allow ion flow through them. They are often called ionotropic receptors. Channels can be "gated" open by many mechanisms including ligand binding, change in membrane potential, lipid interactions, and mechanical stress. Opening a channel to ion flow allows quick passage of information (in this case an electrical signal) into the cell, leading to quick cellular responses. This is an ideal signaling mechanism for neural cells which demand quick responses. We'll show examples of each type of gating mechanism. Before we do, it is helpful to know typical extracellular and intracellular ion concentrations in a mammalian neuron, for example (Table $1$). Table $1$: Extracellular and intracellular ion concentration in a mammalian neuron ion extracellular = [ion]out (mM) intracellular = [ion]in (mM) Na+ 145 mM 5-10 mM K+ 5 mM 140 mM Cl- 110 mM 10 mM Ca2+ (free) 1.2 mM 100 nM When ion channels are opened in neural cell membranes, the direction of favorable thermodynamic flow is down a concentration (chemical potential) gradient but the direction is also affected by the transmembrane potential. Typical resting potentials of neural cells are about -60 to -70 mV (negative inside). If a nonspecific cation channel is gated open, the kinetic barriers to diffusion are relieved and at that moment Na+ ions would flow in due to both the chemical and electrical potential, while K+ ions would flow out but with less driving forces since its efflux is hindered by the negative transmembrane potential. How are such large gradients of these ions formed? We'll answer that in the next section on active transport. Pentameric ligand-gated ion channels (pLGICs) These channels play a key role in neuronal signaling. They are ligand-gated channels. In neural systems, the ligands are neurotransmitters. All are comprised of five monomers, which together form the functional channel with the pore formed in the center of the pentameric structure. The subunits can be identical (homopentamer) or different (heteropentamer). All have "Cys-loop" motifs so they have been called Cys-loop receptors as well. Examples include the mammalian nicotinic acetylcholine, serotonin (5-HT), γ-aminobutyric (GABA), glycine, and glutamate receptors. Figure $1$ shows the generic structure of the pLGICs The monomeric structure is shown on the left. Each contains four transmembrane helices (TM1-4). A top-down view of the pentameric structure is shown to the right. The pore surface forms at the interface of the central TM2 helices. The ligand (neurotransmitter) binds to the extracellular domain with contributions from all the subunits. On ligand binding, TM2 and TM3 rearrange to allow the formation of a transient pore and passive diffusion of specific ions. pLGICs are incredibly interesting and pharmacologically relevant. In general, they have two different types of binding sites. Orthosteric sites bind ligands in the extracellular domains. When bound, a conformational change leads to the rearrangement of helices opening the pore. The natural ligand is also called the agonist as it promotes the function (either neuron excitation or inhibition) of the ion channel. The binding of the natural ligand/agonist opens up the channel to ion flow. This can lead to activation or excitation of the neural cell if positive cations flow into the cell, which depolarizes the cell as the transmembrane potential becomes more positive. Neurotransmitters that lead to this response are excitatory. Alternatively, the binding of inhibitor neurotransmitters in the orthosteric site can lead to inhibition of the neural cell activation if the channel is a ligand-gated anion channel. This hyperpolarizes (makes the transmembrane potential more negative), leading to inhibition of neural cell activation. Inhibitors or antagonists of channel function, whose structure typically resembles at least somewhat the structure of the endogenous ligand or agonist, also bind to the orthosteric site. Allosteric sites are distal to the orthosteric site. Ligands that bind to allosteric sites also lead to conformational changes that either augment or diminish the effect of normal ligand/agonist binding by modulating ion flow through the pore. pLGICs interact with analgesics and anesthetics which makes them even more interesting. Figure $2$ shows two excitatory pLGICs, the 5-hydroxytryptamine (5HT) (left) and nicotinic acetylcholine (right) receptors. The ligand binds in an orthosteric site in the extracellular domain (ECD), which is composed mostly of beta secondary structure. Allosteric sites are often found in the transmembrane domain. Likewise, Figure $3$ shows orthosteric binding sites for GABA and glycine, as well as the binding of modulators that can bind in the TMD or, in the case of benzodiazepines in the ECD as well. How do inhalational aesthetics work? The actual mechanisms of how anesthetics work are still unclear. These fascinating molecules can alter function in a variety of organisms including bacteria, yeast, worms, flies, and plants, as well as animals. Of course, their effect on consciousness appears to be found only in animals. Their selective "turning off" of a function (consciousness) of an entire organ (the brain) is stunning! One theory suggests that they exert their effects through bulk changes in the lipid bilayer, as the potency of anesthetics is generally related to their hydrophobicity. Most are nonpolar and have long been known to work on membranes, presumably altering ion flow through neural membrane ion channels. Typical inhalational and intravenous anesthetics are shown in Figure $3$, along with their date of first use. Figure $3$: Structure of common inhalational and intravenous anesthetics. Adapted from Eur J Anaesthesiol. 2009 Oct; 26(10): 807–820. doi: 10.1097/EJA.0b013e32832d6b0f A very robust correlation is found between the minimum alveolar concentration (MAC, in atmospheres) of inhalational anesthetics and their partition coefficient into olive oil (a measure of their hydrophobicity). This is illustrated in the Meyer-Overton plot shown in Figure $3$. Figure $3$: The Meyer-Overton plot of minimum alveolar concentrations (in atmospheres) vs partition coefficient into olive oil for inhalation anesthetics. https://commons.wikimedia.org/wiki/F...orrelation.png The MAC is the concentration of inhaled anesthetic within the alveoli at which 50% of people do not move in response to a surgical stimulus (i.e. it is much like an IC50 for receptor inhibition). What's so fascinating is the range of molecular species, including N2 and most Noble gases, that can act as anesthetics. Consider N2, a nonpolar and nonreactive molecule, which comprises 80% of the atmosphere. From the graph, it is evident that it takes high pressure for it to exert its anesthetic properties. That can occur when scuba diving using regular air in tanks. Divers can experience nitrogen narcosis (also called depth intoxication or rapture of the deep) when using just compressed air. Hence a mixture of 21% oxygen, 35% helium, and 44% nitrogen is often used. Nitric oxide is used by dentists to alter consciousness and pain perception but does not cause general anesthesia except in some who can be hypersensitive to its use. Additional studies suggest protein:anesthetic interactions are important. For example, the activity of the water-soluble protein luciferase is affected by them. Some molecules (like dichlorohexafluorocyclobutane), expected to have anesthetic properties based on their hydrophobicity, don't. The (S) enantiomer of isoflurane is 50% more potent than the (R) enantiomer in rats, which is hard to explain based on nonspecific partitioning into a bilayer. Most modern theories suggest that they more directly affect specific target proteins and their proximal interacting lipids in neuromembrane bilayers. The main targets of anesthetics appear to be pLGICs. Anesthetics reduce neuron excitability and firing. Hence you could hypothesize that they inhibit excitatory pLGICs (such as the 5HT and acetylcholine receptors) and/or activate inhibitory ones such as the GABA and glycine receptors. pLGICs are pharmacological targets of many general anesthetics. However, anesthetic inhibition of certain GABA channels and potentiation of nicotinic acetylcholine channels have also been observed. Recent elegant studies have shown that the inhaled anesthetic chloroform and isoflurane affect K+ ion flow through the potassium channel subfamily K member 2, known also as the outward rectifying potassium channel protein TREK-1. The channel converts between a voltage-insensitive potassium leak channel and a phosphorylated voltage-dependent outward rectifying potassium channel. It doesn't affect the channel protein directly but indirectly through alteration in the local membrane which affects the location of phospholipase D2 (PLD2), a protein anchored to the membrane by covalent attachment of palmitic acid. PLD2 hydrolyzes phosphatidylcholine, with a positively charged choline head group, to choline and phosphatidic acid, with a negatively charged phosphate head group The effect of these general anesthetics appears to be on lipid rafts in neural membranes. Lipid rafts are enriched in cholesterol and saturated lipids, especially sphingomyelins such as monosialotetrahexosylganglioside1 [GM1]. Rafts are especially important in the brain where cholesterol can reach up to 45% of plasma membrane lipids. Typical rafts are about 100 nm in diameter. In the presence of hydrophobic anesthetics, the rafts become larger and more dispersed, as the anesthetic partitions into them. Membrane proteins also partition into rafts. One such protein is phospholipase D2 (PLD2), which is targeted to the inner leaflet rafts by post-translation palmitoylation. In the presence of general anesthetics, the PLD1 laterally translates away from the disrupted and enlarged lipid raft and binds to a disordered C-terminal region of the TREK-1 protein. This localizes the PLD2 and helps activate it to produce high local concentrations of phosphatidic acid with its negatively charged head group. That group interacts with a positive region of the TREK-1 protein, inducing a conformational change in the channel which opens it to K+ efflux. In effect, PLD2 activates TREK-1 through the local formation of phosphatidic acids. The opening of the channel hyperpolarizes the cell membrane (making the inside more negative, and inhibiting neural activity (a hallmark of anesthesia). These concerted actions are shown in Figure $\PageIndex{a}$ below. Deletion of the TREK-1 gene decreases the effect of the anesthetics. If a catalytically inactive mutant of PLD2 (K758R) is overexpressed, all effects of chloroform were eliminated. We will now explore two pLGIC, the eukaryotic nicotinic acetylcholine channel, and a prokaryotic analog, GLIC. Nicotinic acetylcholine channel (6cnj) One very interesting channel is the one involved in nicotine addiction. It binds nicotine (an exogenous alkaloid) and the normal endogenous neurotransmitter, acetylcholine. Both compete for the same orthosteric binding site. Since the binding of nicotine gates open the channel, nicotine acts as an agonist. The similarities in their structures are illustrated in Figure $4$. The membrane protein is a ligand (acetylcholine)-gated (open-close) positive ion (Na+ or K+) channel, involved in fast neural communication (such as at the neuromuscular junction). The quaternary structure of the pentameric receptor consists of two α4 and three β2 subunits - (α4)2(β2)3. This isoform is the most abundant in the human brain and the one involved in nicotine addiction. The iCn3D model (6CNJ) below has two bound nicotines (spacefill) in the extracellular domain and one Na+ ion (spacefill) in the transmembrane domain containing the pore. The Na+ or K+ ions flow across the membrane down a concentration gradient in a thermodynamically favored process. Figure $5$ shows an interactive iCn3D model of the nicotinic acetylcholine channel with two bound nicotines (spacefill) in the extracellular domain and one Na+ ion (spacefill) in the transmembrane domain containing the pore (6CNJ). (long load time) The Na+ or K+ ions flow across the membrane down a concentration gradient in a thermodynamically favored process. The gold, blue, and brown β2 subunits are glycosylated on Asn 143 and are shown with a Man (β4) GlcNAc (β4) GlcNAc N-linked oligosaccharide. Nicotine is bound between two alpha-beta interfaces. One is shown between the green (alpha) and gold (beta) subunits and the other is between the magenta (alpha) and blue (beta) subunits. GLIC: A prokaryotic pLGIC This protein is a proton-gated cation channel with specificity for both Na+ and K+, which diffuse down their electrochemical gradients. In a sense, H+s in the extracellular side act as "ligands" as the channel is opened with increasing H+ concentration (decreasing pH) on the outside of prokaryotic cells. The protein is homologous to eukaryotic pLGICs. Structures of the protein from Gloeobacter violaceus bound to propofol, an anesthetic, are known. GLICs also interact with ethanol and barbiturates as well. Hence they serve as models to elucidate the binding and effects of anesthetics. In contrast to eukaryotic pLGICs, the "ligand - H+" does not bind in the orthosteric site in the extracellular domain occupied by traditional ligands. Rather changes in protonation states of key proton acceptors and donors in the protein lead to conformational changes analogous to those found on binding ligands to orthosteric sites on classical pLGICs. The external pH associated with half-maximal inward current, pH50 is approximately 5.1 ± 0.2. Evidence suggests that when pH is lowered from 7 to 4, Glu 35 (distant from the orthosteric site), with a pKa - 5.8, becomes protonated. It connects through other H+ acceptors and donors in the open form through a hydrogen bond network. These include two triads of amino acids found at the interface between the extracellular (ECD) and transmembrane (TMD) domains. R192-D122-D32 comprise a conserved "electrostatic triad". The second is Y197-Y119-K248. The network allows bridging of the effects starting with Glu 35 in the ECD into the transmembrane region where allosteric effectors usually interact with the protein. Figure $6$ shows an interactive iCn3D model of the open form of GLIC (3P50) with bound propofol (long load time) Orient the iCn3D model below with the extracellular domain (mostly beta structure) at the top and the transmembrane domain (alpha-helical) at the bottom. Key molecular players involved in the interactions described above, from the top down: • Glu 35 (stick, color CPK) • R192-D122-D32 electrostatic triad (sphere, CPK color) • Y197-Y119-K248 triad (stick, color magenta) • propofol (sphere, color CPK) Propofol and another anesthetic, desflurane, bind at the same site localized in the upper part of the transmembrane domain of each of the five subunits. The model below shows the mostly nonpolar (induced dipole-induced dipole) interactions between one bound propofol and side chains in the TMD. Also shown is an interaction between phosphatidylcholine and propofol. Figure $7$ shows an interactive iCn3D model showing the mostly nonpolar (induced dipole-induced dipole) interactions between one bound propofol and side chains in the TMD. Also shown is an interaction between phosphatidylcholine and propofol. (long load time) Voltage-Gated Ion Channels In contrast to pentameric ligand-gated ion channels, which require 5 monomeric subunits to aggregate into a quaternary structure to form a pentameric pore, voltage-gated ion channels can form a channel from an aggregate of monomeric proteins, each containing a single 6 transmembrane helical unit, or from a longer polypeptide containing multiple repeating 6 transmembrane helical units. Figure $8$ shows a cartoon of a common voltage-gated K+ channel. The monomer (top), denoted as the α subunit, contains a transmembrane domain containing 6 helix segments. Four of these monomers aggregate to form the actual homo- or heterotetrameric channel (bottom). The genes for the Kv channel family, which facilitate K+ diffusion across the membrane, encode α subunits of approximately 500 amino acids and a molecular weight of about 57,000. Four of these α subunits come together in the membrane to form the functional channel, a tetramer of α subunits, which together make one central pore. The α subunit can form homo- or heterotetramers since there are different α subunit encoding genes. In addition, the functional channel has smaller, regulatory β subunits as well. A cartoon structure of a typical voltage-gated Na+ channel is shown in Figure $9$. It is a single polypeptide (α) chain that contains 4 sequential repeats of the 6 transmembrane helical segments (I-IV). The functional channel (bottom) has just one polypeptide chain. The Na+ channel has a molecular weight of around 229K and about 2000 amino acids (each 4x that of the K+ channel α subunit). It is glycosylated and subjected to multiple post-translational modifications. Usually, the protein in the central nervous system is a complex of the α subunit, and small additional regulatory β subunits, which modify the kinetics and voltage-dependency of the α subunit channel. Segment (helix) 4 of each of the four repeat units illustrated above is the conserved "voltage" sensor. It contains multiple, charged amino acids whose disposition changes with changes in the transmembrane potential, allowing conformational changes in the protein and gating of ion flow. Each of the 4 repeating units above also contains an extracellular P-loop (colored purple in segment I in purple) connecting helix 5 and helix 6. We will focus on K+ channels with some additional information on the Na+ channel below. K+ Permeation through Kv1.2 Channel Voltage-dependent potassium channels (Kv) have 4 subunits and can be homo- or heterotetramers. They allow the voltage-gated flow of potassium ions through the membrane. Several obvious questions should arise. How can they be selective for K+ ions? That is, how can they allow the larger K+ ion to passively flow through and not the smaller Na+ ions? Secondly, how can a change in the transmembrane potential cause the channel to open or close? That question boils down to how a change in transmembrane potential can change the conformation of proteins. We'll show several iCn3D models of this protein. Figure $10$ shows a simplified view of the rat Kv1.2 channel (3lut) from top and side views (without parts of the cytoplasmic domain), showing each of the 4 identical monomeric subunits in a different color. Each monomer has S1-S6 transmembrane segments. The protein is found in the brain and central neuron plasma membranes and also in the cardiovascular system. K+ ions diffuse through the center from the extracellular to intracellular side down a concentration gradient but potentially against the transmembrane potential. The four monomers in the homotetramer form one central pore. Figure $11$ shows an interactive iCn3D model showing the rat Kv1.2 channel (3lut) described in Figure $10$. Figure $12$ shows an interactive iCn3D model showing the S4 voltage sensor helix in each monomer of the Kv1.2 potassium channel (3lut) . Figure $13$ shows a more detailed structure of the Kv1.2 potassium channel (3lut). The four monomers, which pack together to form the tetramer, are shown in light and dark grey to allow key residues highlighted in color to stand out. The details are explained below. The left images show the channel from the side (top left view) and top (bottom left view). The right images show just one of the monomers with different side chains highlighted. Figure $14$ shows an interactive iCn3D model detailing key residues in the workings of the Kv1.2 potassium channel (3lut) K+ selectivity - All potassium ion channels, even if not voltage-gated, solve the selectivity dilemma in a similar way. All have in the narrowest part of the pore in the center of the channel this consensus sequence - Thr-Thr-Val-Gly-Try-Gly (TTVGYG) - which is found in the P-loop. These are shown in gold and brown colors in the figure above. The -OHs in the selectivity filter can interact with a dehydrated K ion but not with a dehydrated Na ion, which can not approach close enough to form significant interactions. Surrounding the filter are twelve aromatic amino acids which constrain the size of the pore opening. The interactions of the filter O's with the K ion make up for the energetically disfavored dehydration of the ion. The filter contains K+ ions which repel each other, assisting in the vectorial discharge of the ions through the membrane. These ions must form weak interactions with the selectivity filter. The actual pore is mostly hydrophobic, which facilitates ion flow through the membrane. Figure $15$: below shows a closeup of the selectivity filter. Four Thr 374s (second Thr in the selectivity filter sequence of TTVGYG) from the four different monomers in the channel are clearly shown interacting with the top K+ ion (gray sphere). Figure $16$ shows an interactive iCn3D model detailing key residues in the K+ selectivity filter of the Kv1.2 potassium channel (3lut). Hover over the residues to identify them. Different voltage-gated ion channels alter ion selectivity through changes in these amino acids in the P-loop, as illustrated in Table $2$ below. As channels lose specificity of K+, they gain specificity for Na+ and Ca2+. Red highlights denote conserved residues and yellow residues that are chemically similar. sequence specificity TVGYG strong K+ channels CIGYG weak K+, HCN channels TVGDG TRP channels STFEG ionotropic glutamate receptors LCGEW strong Ca2+ voltage-gated channels Table $2$: P-loop specificity side chains in voltage-gated ion channels Voltage gating - Helix S4 in each monomer in the transmembrane domain of the complex is the voltage sensor. The sequence of this helix is LAILRVIRLVRVFRIFKLSRH. Note the arginines and lysine highlighted in blue. They repeat every 3 amino acids. The voltage-sensor domain must be shielded from the nonpolar acyl of the bilayer. Four conserved Arg residues on S4, part of the voltage-sensor domains, are shielded from the lipids and coupled to an amphiphilic helix running parallel to the plane of the membrane. The arginines move under the influences of forces arising from changes in the membrane's electric field initiated by ion movement through other ion channels in the membrane. Mechanical work is done by the electric field on the voltage sensor as the charged Arg residues are moved through the electric field. The movement is coupled through the amphiphilic helix to the pore which changes conformation. In turn, the S4 and coupled S5 helices of the voltage sensor do mechanical work on the pore by altering its conformation to open/close the pore, specifically at the activation gate of the pore. This seems quite similar to how iron movement into the heme plane in hemoglobin on oxygenation pulls the proximal His on the F8 helix which then transmits a conformation change to other helices in the subunit, leading to cooperative conformational changes in the tightly packed protein. About 12 charges move across the transmembrane potential field. Channels, once open, must be inactivated. In the case of the voltage-gated potassium channel, inactivation occurs when the amino-terminal cytoplasmic domain binds to the potassium pore on the cytoplasmic side, in interaction likened to the binding of a "ball on a chain" (the cytoplasmic domain) to the pore opening. The chain acts to tether the ball domain so it may swing to bind to the pore opening. The ball domain contains both positively charged and hydrophobic regions. Where is the ball domain in the absence of inhibition? Recent studies (Oliver et al.) have shown that a positive domain can bind to proximal phosphatidyinositol 4,5-bisphosphate (PIP2) lipids on the inner leaflet of the membrane bilayer. When so bound, inactivation of the channel is prevented. As you will see in the next section, PIP2 can also be cleaved to form diacylglycerol and inositol 1,4,5-trisphosphate when cells are activated by external factors (hormones, growth factors, etc) in the process of signal transduction. Figure $17$ shows a molecular dynamics simulation showing K+ interaction with the channel lining and the "knock-on" mechanism showing how an incoming K+ ion can repel a K+ ion in the pore through the channel. Figure $17$: Molecular dynamics simulation of K+ movement through a channel Voltage-gated sodium channels (NaV) The eukaryotic voltage-gated sodium channels (NaV) allow inward movement of Na+ ions which depolarizes a neuron and sets off an action potential in nerve and muscle cells. (For more information on neuron signaling, see Chapter 28.9).   The NaV has an alpha subunit that forms the pore plus beta subunits that associate with it and modulate its activity. Nine eukaryotic isoforms exist.  NaV has 4 domains, I-IV,  each containing segments 1-6. Each of the S1-S4 segments forms a voltage-sensitive domain and each of the S5 and S6s form the pore.  In contrast to eukaryotic NaVs which have a single chain, bacterial NaVs contain 4 identical subunits.  Post-translation modification of the alpha subunit can regulate its activity. As important as it is to initiate Na+ influx to trigger neuron firing, it is equally important to turn it off to control neural signaling.  The inward Na+ current is stopped by a fast inactivation that occurs within a few milliseconds. Many neurotoxins bind to the NaV and regulate its activity.  Key examples are the α-scorpion and sea anemone toxins which both inhibit the fast inactivation of the NaV leading to prolonged or sequential action potentials.  These toxins binds to the voltage sensor in domain IV which is key for the fast inactivation in the absence of the toxin. The S4 helical segments in each domain are the key voltage sensors.  Each of the S4 segments has 4-6 positively charged Arg and Lys side chains. On depolarization of the cell (when the inside becomes less negative and more positive), this helix move "up" from the cytoplasmic side (which is increasingly more positive) which opens voltage-gated channel.   Specificity for the small Na+ ion (over the K+ and Ca2+ ions) is determined mainly by 4 amino acids, DEKA (Asp-400, Glu-755, Lys-1237, and Ala-1529) found in P loops of domain I-IV, respectively. This selectivity filter is conserved. Figure $18$ below shows a cartoon of the NaV with two associated, regulatory beta chains. Figure $18$: Cartoon of eukaryotic voltage-gated sodium channel (Nav) with two associated, regulatory beta chains.   https://www.guidetopharmacology.org/...rd?familyId=82. CC BY-SA 4.0 Note the sites for posttranslational modification by phosphorylation and drug interactions.  Cylinders represent probable α-helical segments S1-S6. Bold lines represent the polypeptide chains of the selectivity filter and tetrodotoxin binding site; The yellow S4 segments are the voltage sensors.  The "h" in the blue circle is in the inactivation gate loop.  Blue circles are sites implicated in forming the inactivation gate receptor. Sites of binding of α- and β-scorpion toxins (ScTX) and a site of interaction between α and β1 subunits are also shown. Tetrodotoxin is a specific blocker of the pore of sodium channels, whereas the α- and β-scorpion toxins block fast inactivation and enhance activation, respectively, and thereby generate persistent sodium current that causes depolarization block of nerve conduction. Table $3$ shows the different types of neurotoxin receptors sites found in NaVs Neurotoxin Receptor Site # Toxin or Drug  Domains 1 Tetrodotoxin IS2–S6, IIS2–S6 Saxitoxin IIIS2–S6, IVS2–S6 µ-Conotoxin 2 Veratridine IS6, IVS6 Batrachotoxin Grayanotoxin 3 α-Scorpion toxins IS5–IS6, IVS3–S4 Sea anemone toxins IVS5–S6 4 β-Scorpion toxins IIS1–S2, IIS3–S4 5 Brevetoxins IS6, IVS5 Ciguatoxins 6 δ-Conotoxins IVS3–S4 local anesthetic drug sites Local anesthetic drugs IS6, IIIS6, IVS6 Antiarrhythmic drugs Antiepileptic drugs Table $3$: Neurotoxin receptor sites in NaVs.  https://www.guidetopharmacology.org/...rd?familyId=82. CC BY-SA 4.0 A view of the effective pore in the bacterial Nav is shown below in Figure $19$. Figure $19$: Structure of the bacterial sodium channel NavAb pore B. Architecture of the NavAb pore. Glu177 side-chains in the P loop are shown in purple.  The pore volume is shown in grey. The P and P2 alpha helices that form the scaffold for the selectivity filter and outer vestibule are shown in green and red, respectively. https://www.guidetopharmacology.org/...rd?familyId=82. CC BY-SA 4.0 As mentioned above, the bacteria NaVs have 4 monomeric subunits.  In contrast to the K+ channel, which requires the K+ ions to be dehydrated to make sufficient interactions with the pore and to pass through, the Na+ ions need to be hydrated. Figure $20$ shows an interactive iCn3D model of the A and B chains of the bacterial NaV voltage-gated sodium channel pore and C-terminal domain (5BZB) Figure $20$:  A and B chains of bacterial NavMs voltage-gated sodium channel pore and C-terminal domain (5BZB) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...vRmDqmDHGsYLh8 One subunit is shown in a transparent cyan surface and the second one is shown in magenta.  Three Na+ ions are shown (gray spheres).  Eight water molecules are shown interacting with them in the pore.  The other two subunits are not shown for clarity. The voltage-gated sodium channel has three major conformational states: • a basal closed state found at resting cell potentials in which the pore of Na+ is occluded by an activation gate • an open state found when a depolarizing potential is reached in the cell • an inactivated state formed within 10 ms of opening of the channel when the inactivation gate with a a Ile-Phe-Met (IFM) sequence motif, found in the intracellular linker between domain III and IV (near the cytoplasmic face of the receptor) closes off the pore to further Na+ entry. The protein converts back to the closed state when the transmembrane potential is restored to its initial value (around -70 mV) and the positively charged S4 segments move back towards the cytoplasmic face. A cartoon of the three-state model for the Na+ channels (as other voltgage-gates ion channels in general) is shown in Figure $21$ below. Figure $21$: Three-state model for the voltage-gated ion channels.  Hinard, Valerie & Britan, A & Rougier, Jean-Sébastien & Bairoch, Amos & Abriel, Hugues & Gaudet, Pascale. (2016). ICEPO: The ion channel electrophysiology ontology. Database : the journal of biological databases and curation. 2016. 10.1093/database/baw017. Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0/) The YouTube video by Pete Meighan below shows an incredibly clear description of the three conformational states of the channel and the conversion from closed to open to inactivated states. To understand these conformations states, we need to look a the structure of the protein in greater detail.  Figure $22$ below shows multiple representations of the voltage-gated sodium channel. The pore domain PD is formed from helix segments 4 and 5 on each of the IV domains of the protein. The voltage sensor domain is formed from the S1-S4 segments, of which segment S4, containing multiple positively-charged Arg and Lys, is key. Figure $22$: Structure features of the voltage-gated sodium channel.  Dongol, Y.; C. Cardoso, F.; Lewis, R.J. Spider Knottin Pharmacology at Voltage-Gated Sodium Channels and Their Potential to Modulate Pain Pathways. Toxins 201911, 626. https://doi.org/10.3390/toxins11110626.  Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/) Panel (A) shows a schematic representation of the α-subunit of voltage-gated sodium (NaV) channel. Four non-identical domains (DI–DIV) feature six neurotoxin receptor sites (Sites 1–6) and key residues contributing to the outer Na+ ion selectivity filter (EEDD) and inner selectivity filter (DEKA). The connecting S5–S6 linker is called P-loop (P) which together with S5 and S6 segments from each domain contributes to the formation of the Na+ ion selective channel pore.  Sites 1-6 (colored purple, green, cyan, magenta, etc) are sites where inhibitors such as toxins bind. Panel (B) shows the structure of the NaV1.7 channel (PDB 6J8G). Four voltage sensing domains (VSDs), DI (yellow), DII (blue), DIII (green), and DIV (orange), are shown with their corresponding pore-forming segments (S5 and S6) arranged to form the pore domain (PD) selective to Na+ ions. The P-loop that contributes to forming the inner selectivity filter is colored in red spheres (DEKA) and outer selectivity filter (EEDD) is colored in purple. The S6 segments of all the four domains contribute to form the intracellular region of the pore. Site 3 (cyan) and Site 4 (pink) are the major binding sites for spider knottins (neurotoxins). The β1 and β2 subunits which interact with DIII and DI, respectively, are highlighted in beige color. Panel (C) shows a schematic of the three main conformational states of the protein which control gating of NaV channels. At polarized potentials, the DI–DIV S4 segments are drawn towards the intracellular side due to the positive gating charges to render the closed conformation (down state). Upon depolarization, the forces holding the down state are relieved and DI–DIII S4 segments are rapidly released extracellularly to open the S6 channel gate in the open conformation (up state). Note the movement of the S4 helix with its positive charged toward the extracellular side of the membrane. The DIV S4 moves up slowly compared to DI–DIII S4 and drives the fast inactivation, where the channel is occluded intracellularly by the Ile, Phe, and Met (IFM) motif. After cell repolarization, the channel returns to a closed (resting) state. Figure $23$ shows an interactive iCn3D model of a ternary complex of human Nav1.2 with the beta2 regulatory subunit and conotoxin IIIA (6J8E) Figure $23$: Human sodium channel Nav1.2-beta2-KIIIA ternary complex (6J8E) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?aYxomdUqZHqbMmE5A Domains I-IV are shown in gray, yellow, green and cyan, respectively. The separate beta-2 regulatory subunit is shown in magenta. The positive side chains in each of the S4 segments of the four domains are shown as red sticks.  The side chains of the inner 4 amino acids (DEKA) comprising the selectivity filter are shown in spacefill, CPK colors, and labeled.  The brown peptide cartoon on the extracellular side (red sphere layer for outer leaflet) is the µ-conotoxin KIIIA.  The spacefill molecule in the pore near the blue sphere layer representing the inner leaflet is the neurotoxin veratridine (VTD), which inhibits channel inactivation and lengthens the action potential with possibly fatal consequences.  A single Na+ ion is shown at the top of the pore as an orange sphere labeled Na.  The µ-conotoxin blocks the pore.  The Ile1488-Phe1489-Met1490 (IFM) motif, found in the intracellular linker between domain III and IV (near the cytoplasmic face of the receptor, responsible for the fast inactivation, are shown in gray spheres and labeled with single-letter codes. The selectivity filter DEKA is different from the selectivity filter in another sodium channel (NavAb) which has 4 glutamate.  Asp and Ala line the wall of the filter region and Glu and Lys can attract and release, respectively, the Na+ ion. For fast inactivation, the IFM motif must interact with an"inactivation gate receptor" within the protein itself. Likely candidates for this are short intracelllar loops connecting all the S4 and S5 segments.  This receptor appears to contain 3 amino acids, F1651, L1660 and N1662. Figure $24$ shows another interactive iCn3D model of a ternary complex of human Nav1.2 with the beta2 regulatory subunit and conotoxin IIIA (6J8E) highlighting just the DEKA selectivity filter, the IFM motif, and its receptor inactivation gate F1651, L1660 and N1662. Figure $24$: Human sodium channel Nav1.2-beta2-KIIIA ternary complex highlighting just the DEKA selectivity filter and the IFM and its receptor inactivation  gate F1651, L1660 and N1662 (6J8E).  (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?tq3kXbT44GeCCUcY7 They are all shown in spacefill and labeled.  Orient the molecule with the cytoplasmic domain at the top.  The leaflets of the bilayer are omitted for clarity but the cytoplasmic pore inhibitor is still shown in sticks. Figure $25$ below shows a closeup of the bound Na+ ion. Figure $25$: Sodium ion in the SF of the PDB ID 6J8E Nav1.2 cryo-EM structure. The pore blocker μ-conotoxin KIIIA is not shown. Alberini et al. J. Chem. Theory Comput. 2023, 19, 10, 2953–2972. April 28, 2023. https://doi.org/10.1021/acs.jctc.2c00990. Creative Commons. Structures of the closed state, the open state, and the inactivated state of NaV1.5 are now known.  Key regions are shown in Figure $26$ below.  The open state was the hardest to solve since it closes in milliseconds to form the inactive state as described above.  Mutations in the Ile-Phe-Met (IFM) motif to QQQ prevented inactivation.  This would ordinarily be deleterious to an organism but in the presence of a small molecule inhibitor, propafenone, it was possible to isolate this state. Figure $26$:  Closed, open, and inactivated conformations of the activation gate and the locations of arrhythmia mutations.  Daohua Jiang, Richard Banh, Tamer M. Gamal El-Din, Lige Tonggu, Michael J. Lenaeus, Régis Pomès, Ning Zheng, William A. Catterall.  Open-state structure and pore gating mechanism of the cardiac sodium channel.  Cell, Volume 184, Issue 20, 2021, 5151-5162.e11, ISSN 0092-8674,. https://doi.org/10.1016/j.cell.2021.08.021. Reprinted with permission from Elsevier.  May not be sublicensed, assigned, or transferred to any other person without publisher's written permission. Panel (A) shows the closed activation gate of NaV1.5 generated by MODELER based on the resting-state structure of NaVAb (PDB: 6P6W), sealed by a square of hydrophobic side chains of hydrophobic residues V413, L941, I1471, and I1773 (spacefill, black) that in the closed state completely seal off the cytoplasmic opening in the pore. These ring of amino acid side chains come together on conformational changes resulting from the engagement of the IFM motif with its internal receptor. Panel (B) shows the open activation gate of NaV1.5/QQQ triple mutation. Red arrows indicate the directions of movement of S6 segments compared to the resting state. Panel (C) shows the partially open but nonconductive activation gate of rNaV1.5C in the inactivated state. Red arrows indicate the directions of movement of the S6 segments compared to the open state. Panels (D–F) show structures from (A) to (C) are shown in space-filling surface representation with hydrated Na+ placed in the central cavity behind the activation gate. Red and green spheres represent water and Na+, respectively. van der Waals distances measured across the orifice of the activation gate are 4.3 Å (DI-DIII) × 2.8 Å (DII-DIV) for the resting/closed state, 6.9 Å (DI-DIII) × 5.0 Å (DII-DIV) for the inactivated state, and 7.3 Å (DI-DIII) × 8.2 Å (DII-DIV) for the NaV1.5/QQQ open-state structure. Panels (G–I) show structures from (A) to (C) with the locations of arrhythmia mutations causing LQT-3 overlaid as green spheres The spacefill models in D-E clearly show that the cytoplasmic opening is "open" in the open state E (obviously), and occluded to increasing degrees in the inactive state (F) and closed state D. Finally, figure $27$ shows an interactive iCn3D model of the electrostatic surface potential of the rat Nav1.5 channel (6UZ3) Figure $27$: Electrostatic surface potential of the rat Nav1.5 channel (6UZ3) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...UZdQeUUPtYGgR6 The red indicates negative potential and the blue positive potential.  The white is neutral.  Note that the bilayer representations are essentially neutral with respect to their effect on the overall surface potential. Tilt the model to view the top and bottom entrance to the pore and the overall negative charge density expected in the net attraction of the positive Na+ ions. Lipid-Gated Ion Channels Membrane receptors are in the lipid bilayer, so it should not be too surprising that specific lipids might bind and trigger conformational changes in the receptor, mediating a specific biological activity. The specific lipid might form during an upstream event, then bind to the receptor, triggering function in a process of signal transduction, which we will explore in the next chapter. Let's look at receptors gated by binding of the lipid phosphatidylinositol bisphosphate (PIP2). It is found in small concentrations in membranes and can be cleaved in by phospholipase C to form a small polar intracellular signaling molecule, IP3 (discussed in Chapter 12). PIP2 can also recruit proteins to the membrane and participate in signaling events that way. For this section, we will discuss how it binds to a K+ ion channel protein and open so K+ can flow into cells (the opposite direction of the normal efflux), which helps control transmembrane potential. Kir2.2 -Inward rectifier potassium channel Kir2.2 The names of this channel can best be understood if you understand the meaning of the word rectify. The verb has several meanings but for us, the best definition is to correct. What does this channel correct? When is it functional? We'll explore the details more in the next chapter we first a better understanding of what happens to the actual Na+ and K+ ion concentrations outside and inside the cell during neural activation. How much do they change? We need this to understand the driving force for the Kir2.2 channels that move K+ into the cell, which is opposite the usual direction. From that sense, it is "rectifying" the K+ concentrations. How much do the actual concentrations of Na+ and K+ ions change when a neuron is excited? When a neural cell is activated or fires, the cell membrane potential goes from a resting potential of around -60 to -70 mV (inside negative ) to a more positive potential as Na+ ions enter the cell through voltage-gated Na+ channels (after an early neurotransmitter-gated ion channel is open after ligand binding) with the ions flowing down a chemical and electric potential gradient. At a certain membrane potential (about +30 mV), K+ channels open allowing an efflux of K+, also down both a chemical and electric potential gradient, returning the cell potential close to its equilibrium value potential of around -60 to -70 mV (inside negative). But how much do the actual K+ and Na+ ion concentrations change in this process? The somewhat counterintuitive answer is hardly any at all! We need to understand how the membrane acts as a capacitor first. The charge Q on a surface of a plate or side of a membrane is proportional to the voltage across the plate or membrane. Figure $28$ shows how a membrane with a transmembrane potential acts as a capacitor. The dielectric medium in the capacitor will determine how quickly the charges on the plate will dissipate. When the medium is an insulator, resistant to charge flow, the plates remained charged longer. The same is true for the membrane. The hydrophobic bilayers act as an insulator and resist the discharge of the membrane potential. The bilayer offers high resistance (low conductance) to charge flow. Only when channels are open and the ions become reasonably permeable to flow does the membrane potential change over short periods. The following derivation is adapted. It makes sense that the stored charge (Q) on either side of the membrane is proportional to the membrane voltage. We can write the following simple equations: \begin{array}{l} Q \propto V \ Q=\mathrm{C} V \end{array} where C, the proportionality constant, is the capacitance with units of the Faraday (which you remember from introductory chemistry). Let's normalize this equation for an area of 1 cm2. The measured capacitance of lipid bilayers is about 10-6F/cm2. Let's assume a voltage change from -70 mV to + 30 mV for a total of 0.1 V. Hence Q=\frac{10^{-6} \mathrm{~F}}{\mathrm{~cm}^{2}}(0.1 \mathrm{~V})=\frac{10^{-7} \mathrm{Coul}}{\mathrm{cm}^{2}} Let's change that into the number of elementary charges on the surface of the membrane per μm2 surface area. \left(\frac{10^{-7} \mathrm{Coul}}{1 \mathrm{~cm}^{2}}\right) x\left(\frac{1 \text { ion }}{1.6 \times 10^{-19} \mathrm{Coul}}\right)=\frac{6.25 \times 10^{11} \text { ions }}{\mathrm{cm}^{2}} \times \frac{1 \mathrm{~cm}^{2}}{10^{8} \mu \mathrm{m}^{2}}=\frac{6,250 \text { ions }}{\mu \mathrm{m}^{2}} What does this mean for an ordinary eukaryotic cell? Let's model the cell as a sphere with a diameter of 10 μm. Knowing the equations for the volume (V = (4/3)πr3) and surface area (4πr2), the volume of 10 μm cell is about 524 μm3 and the surface area is 314 μm2. The table below shows the resting ion concentrations in a cell and the actual number of ions in the 524 μm3 volume of the cell (column 3). Using the calculated value of 6250 ions moved/μm2, the total number of K+ and accordingly Na+ ions that move across 314 μm2 of total cell membrane surface area is about 2 million. The results are shown in Table $3$ below. 1. ion 2. [ion]intracellular (mM) 3. # ionsintracellular 4. total ions moved during neuron response 5. change in [ion]intracellular sodium 10 mM 3.2 × 109 Na+ ~ 2,000,000 in on depolarization ~6.3 x 10-2 mM (~0.6% change) potassium 150 mM 4.7 × 1010 K+ ~ 2,000,000 out on repolarization ~6.3 x 10-2 mM (~0.6% change) Table $3$: How many ions move across a membrane \frac{2 \times 10^{6} \mathrm{Na}^{+} x\left(\frac{1 \mathrm{~mol} \mathrm{Na}^{+}}{6.022 \times 10^{23} \mathrm{Na}^{+}}\right)}{524 \mu M^{3} \times\left(\frac{1 \mathrm{~L}}{10^{15} \mu M^{3}}\right)}=6.3 \times 10^{-6} \mathrm{M}=6.3 \times 10^{-3} \mathrm{mM} The actual change in intracellular Na+ ion concentration on excitation is only about 0.6% of the initial [Na+]intracellular on excitation and depolarization of the cell. We can also assume that the K+ ion changes to the same degree in repolarization. So when the Na+ and K+ channels open, the "flood gates" are not opened. Permeability does increases, but this leads to a tiny influx of Na+ ions, which is not sufficient to change the intracellular Na+ concentration. It is, however, enough to change the transmembrane potential! It's a misconception that there are significant changes in ion concentration across the membrane on depolarization and repolarization of the cell. The Goldman-Hodgkins-Katz equation (previous section), shows that the membrane potential is a function of both concentrations and permeability coefficients. Now we can present the inward rectifying potassium channels, which facilitate the movement of K+ into the cell from the outside. This is in the opposite direction of the usual flow of the ions. Since the ions are moving towards higher levels of K+ inside the cell, it would appear this would be an example of either passive diffusion favored only by the electrical potential or a case of active transport. None of these is true. K+ ions diffuse from outside to inside the cell since outward diffusion is substantially blocked by molecules such as polyamines and Mg2+! We don't have to invoke active transport or a violation of the basic rules of thermodynamics. The channels thus display strong inward currents and weak outwards ones. The channels allow large conduction of K+ ions if the membrane potential is more negative compared to the resting K+ ion equilibrium potential but less if more positive, so the net effect is to maintain the resting K+ potential. There are several subfamilies of Kir channel. They are always potentially active (open) except those gated by G-proteins (see next chapter) or by ATP binding. (these are involved in metabolism). Kir activity is regulated by phospholipids and proteins. Now we will explore the Kir 2.2 channel, which is gated-open by PIP2, a membrane lipid, not by changes in the transmembrane potential. Yet by opening this channel and allowing the inward flow of K+ ions, PIP2 is regulating the transmembrane potential. It is the agonist for the Kir 2.2 channel. Instead of having monomeric units with S1-S6 transmembrane helical segments with a voltage sensor (S4) and P loop and S5-S6 selectivity filter, it has just 2 transmembrane helices. Both the N- and C-termini are in the cytoplasm, connected by an extracellular loop (H5) which helps form the prototypical K+ selectivity filter with the consensus sequence T-X-G-Y(F)-G. Similar to the voltage-gated K+ channel, four of these aggregate to form homo- or heterotetramers in the membrane. Figure $29$ illustrates these points. The proteins have large intracellular domains (ICD). On binding of PIP2 to the region between the transmembrane and ICD, which produces a large conformational change, allowing K+influx. The model below shows the R186A mutant tetrameric Kir 2.2 channel (3SPG) with four bound PIP2 analogs containing two short fatty acids (octanol) esterified to the glycerol backbone. There appear to be two binding sites for lipids, a nonspecific site in the TMD and a specific one for PIP2 in the ICD. Figure $30$ shows an interactive iCn3D model of the inward rectifying R186A mutant tetrameric Kir 2.2 channel (3SPG) with four bound PIP2 analogs containing two short fatty acids (octanol) esterified to the glycerol backbone. Hover over the residues to identify them. Figure $31$ shows an animation that shows the monomeric Kir protein morphing from the apo state (without PIP2, 3JYC) to the PIP2 bound state (3SPI). Another PIP2-gated ion channel is the transient receptor protein (TRP) channel. These channels play a role in vascular tone. Smooth muscle cells have TRPC3 and TRPC6 channels, Na+ or Ca2+ channels that cause depolarization, leading to smooth muscle contraction and vasoconstriction. Mechanosensitive channels We saw in the previous chapter that some pores (not channels) are gated open not by voltage, or specific agonists such as neurotransmitters or even specific lipids like PIP2 but by more general changes in membrane bilayers properties (membrane tension, curvature, pressure). Likewise, specific channel proteins can be gated by changes in the physical properties of the bilayer. We will consider one mechanically-active channel, TRAAK. TRAAK - Potassium channel subfamily K member 4 The protein is a K+ ion channel that is not sensitive to voltage changes but is open on the mechanical deformation of the bilayer. The channel can be opened by making the cytoplasm basic, raising the temperature. It is involved in pain sensation and pressure transduction. The TRAAK 4 opening is blocked by the presence of lipid from the inner leaflet which occludes the pore. If the lipid is removed by physical changes, transmembrane helix 4 rotates, which prevents the lipid from blocking the channel, which opens. Further changes in the coupled transmembrane helices 2 and 3 stabilize the opening. Figure $32$ shows an interactive iCn3D model of the TRAAK channel protein (4wff) in the closed state. Hover over the residues to identify them. The K+ ions are aligned in the channel. Decane, a nonpolar molecule, is shown in spacefill and colored cyan. The decane is probably decanoic acid in which the carboxyl group was not defined in the structure due to high flexibility. This suggests that the "decanoic acid" is not tightly bound. It binds through the cytoplasmic side through an opening in the membrane protein. In the open state, transmembrane helix 4 rotates, blocking access to the cavity. Hence lipid binding gates the channel closed.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/11%3A_Biological_Membranes_and_Transport/11.03%3A_Diffusion_Across_a_Membrane_-_Channels.txt

Search Fundamentals of Biochemistry Pores and Pore-Forming Proteins (PFPs) If you form a pore in a cell bilayer, molecules of all sizes could move either way based on their electrochemical potential. They will move from regions of a higher to lower electrochemical potential in a thermodynamically favorable process. Hence movement through pores represents a special case of facilitated diffusion when part of the driving force is not just a concentration gradient but also an electrical potential. Several questions might come to mind. • What proteins are involved in pore formation? • How is the specificity of solute movement through the pore regulated? • What is the mechanism of pore formation? Pore formation can lead to cell death, which is the function of some pore-forming proteins (PFPs) including the toxin Hemolysin E (also known as HlyE, ClyA, SheA) secreted from E. Coli and S. Aureus. Human proteins also form a membrane attack complex (examples include the membrane attack complex-perforin/cholesterol-dependent cytolysin (MACPF/CDC) superfamily and the membrane attack complex (MAC). The MAC consists of an assembly of proteins involved in the complement system (part of the effector branch in the innate immune system), which leads to cell death of Gram-negative bacteria like E. Coli. Figure $1$ illustrates the assembly process of the membrane attack complex and the complexity of interactions required to form a lethal pore in a cell. Complement protein C9 can adopt a soluble form or membrane form which in aggregates form the pore leading to cell death. This is a common feature of PFPs. PFPs could create a pore by altering membrane lipid packing to form a toroid-like hole (Figure $2$) and/or by inserting in a membrane and forming a pore within the protein complex itself. In either mechanism, lipid packing is altered. Biophysical evidence shows some support for the formation of "toroidal pores". Lipid rearrangements in the membrane could lead to a hydrophilic or hydrophobic pore lining as shown in Figure $3$. We started our study of lipid bilayers with pure lipid systems and then added membrane proteins. Let's do the same with pore formation. A common technique to form a pore in pure lipid bilayers and also in cells is electroporation. This is a technique used to move a DNA with a target gene into either a prokaryotic cell (transformation) or eukaryotic cell (transfection) for exogenous gene expression. In the absence of PFPs, this requires the alteration of surface tension by applying an electrical potential. This forms depressions in the membrane, altering the nonpolar acyl chain packing. In the process, small wire-like columns of water appear, which like hydrophobic pores ultimately rearrange into hydrophilic toroidal pores. Figure $4$ shows snapshots of molecular dynamics simulations as a function of the time of pore formation in electroporation. How does DNA pass through the pores in the bilayer? In pure lipid vesicles, it appears to pass through by electrophoresis. Most students are familiar with the electrophoresis of DNA fragments through pores in agarose gels. In actual living cells, small nucleic acids like small interfering RNA (siRNA) and antisense DNA molecules appear to pass through the bilayer by electrophoresis. Large DNAs like plasmids containing a gene for expression bind to the cell and form cell surface aggregates, which appear to be endocytosed into the cell. Electroosmosis, the movement of liquids under the influence of an electric field, also plays a role. Pores - Outer Membrane Factor (OMF) and Voltage-Dependent Anion Channel (VDAC) Now let's consider pores made of PFPs. We have already discussed two types of beta-barrel transmembrane proteins, the outer membrane factor (OMF) of Gram-negative bacteria and the voltage-dependent anion channel (VDAC). Both are examples of proteins called porins with typical beta-barrel topology. VDAC: At low membrane potentials, VDAC (also known as mitochondrial porin), the most abundant protein in the mitochondrial outer membrane, moves metabolites and Ca2+ ions across the outer membrane of mitochondria. VDAC exist in an open state at 0 or very low transmembrane potentials that allows for the transfer of key metabolic anions (pyruvate, oxaloacetate, malate, succinate, ATP, ADP, and Pi, which are involved in metabolism) and Ca2+, and in a closed state (above or below + 30 mV), which is not completely closed as it allows for the transfer of ions with a preference for cations. The closed state does not allow for the transfer of ATP. The transition to the closed state is promoted by tubulin and actin (cytoskeletal proteins), negatively charged lipids such as phosphatidyl ethanolamine and cardiolipin, and also covalent phosphorylation by protein kinases. Bcl2, proteins involved in the regulation of programmed cell death (apoptosis) also interact with VDAC. In contrast to ligand-gated channels which require ligand binding to open the channel, pore complexes are unusually open. Millions of ATPs/second move across the membrane through the open pore but none through the closed pore. Figure $5$ shows an interactive iCn3D model of mouse VDAC1 (4c69) with bound ATP loosely held in the site. An alpha helix partly occludes the central pore of this β-barrel protein. One ATP is bound in the barrel and interacts with Lys 12 and Lys 20 at each end of the cavity-bound helix. The alpha-helix narrows the pore opening and presumably changes orientation in a voltage-sensitive fashion to gate the pore open and closed, hence regulating the conductance of ions through the pore. That the orientation of the charged arginine side chains would be dependent on the transmembrane potential should be somewhat obvious. Aquaporins Aquaporins can move a billion water molecules per second across membranes and exclude ions including protons. Waters proceed in a single file through the pore. Instead of moving waters through, it might simply move "H+ ions" through by the alignment of subsequent H bond donors and acceptors in a "wire" of water molecules. This is prevented by two conserved asparagine residues in the center of the channel, which disrupt water to water hydrogen bond network in the channel waters that could facilitate proton transfer. Instead, the central waters form hydrogen bonds to the central asparagines. This, along with local membrane potentials, causes opposite orientations of the water in different leaflet sides of the membrane, precluding H+ transfer. Here is a movie of a molecular dynamics simulation of water moving through a porin called aquaporin, GlpF, from E. Coli. Science magazine (Tajkhorshid et al., Science Apr 2002, 296:525). Used with permission from the Theoretical and Computational Biophysics Group, the National Institutes of Health (NIH) Resource for Macromolecular Modeling and Bioinformatics, at the Beckman Institute, University of Illinois at Urbana-Champaign. OMF: The outer membrane factor (see previous section) is one member of a class of bacterial porins, which are the most abundant proteins in the outer membrane of Gram-negative bacteria. They are classified as non-specific or specific (with respect to the solute that passes through), or monomeric, dimeric, or trimeric based on their structure. In Gram-negative bacteria, which have two lipid bilayers, the movement of solute from inside to outside includes at least three sets of proteins. Active transport (discussed in the next section) needs an energy source and is used by inner membrane transport proteins, including ATP-binding cassette (ABC)-type, resistance nodulation division (RND)-type and major facilitator superfamily (MFS)-type transporters. These are connected to membrane fusion proteins (MFP) that span the periplasm, which then interacts with at least 21 different types of porins. Molecules with molecular weights greater than 600 generally can not get through the nuclear envelope of Gram-negative bacteria, limiting the size of potential antibiotics, which must enter by passive diffusion. Mechanosensitive ion channels - Mscs (which are pores!) As the name applies, these ion channels (with openings large enough to be called pores) are gated open/closed by mechanical (physical) changes in the properties of the membrane, not extracellular/intracellular ligands or voltage changes. Certain bilayer lipids also activate the Mscs. There are two types, small (MscS) and large (MscL) mechanosensitive ion channels. Changes in local (boundary layer) and nonlocal lipids are involved in the gating of the channel (in the next section we will discuss lipid-gated ion channels). They are found in prokaryotes, archaea ,and eukaryotes. They are also called stretch-gated ion channels. Mcss transduces a physical force (stretching and change in turgor pressure) into an electric signal - a flow of ions across the membrane. Turgor pressure is the internal pressure that "presses" the cytoplasm and cell membrane towards the cell wall in bacteria and plant cells. It arises mostly from the osmotic flow of water into the cell. If bacterial cells are placed in a high salt concentration solution (hypertonic), water flows out of the cell and the cell membrane shrinks to the inside of the volume confined by the cell wall. When placed in a hypotonic solution, water flows into the cell and the cell membrane presses out to the cell wall. The response of these channels is fast, in the millisecond range, which is about as quick as a cellular response can be. Some variants of these channels are called piezochannels, based on the piezoelectric effect that describes how a voltage is produced when some materials are deformed by mechanical stress which causes a redistribution of charges. Bacteria normally have high concentrations of both K+ and negatively charged anions, especially glutamate, which leads to a high turgor pressure from the inward osmotic flow of water. At low external osmolarity, turgor pressure in the cell could be as high as 4 atm. If placed in external solutions of high osmolarity, bacterial cells respond by the increased movement of solutes into the cell. Mscs are particularly important when bacteria are subjected to sudden osmotic shock. If they are placed in pure water, for example, water would flow down a concentration gradient into the cell and cause the cell to swell and lyse, killing the cell. This is done in the lab to prepare almost pure hemoglobin from ruptured red blood cells. The Mscs are opened under these conditions and small species from the cytoplasm flow out, helping to keep the cell viable. Their openings must be regulated to prevent too much outward flow, which would kill the cell. In other organisms they are also involved in touch (stretch), hearing (vibration, sound waves), and responses to gravity. Stimuli to activate them include fluid shear stress (relevant to endothelial cells that line blood vessels), membrane stretch (relevant to skeletal and cardiac muscle cells), or even indentation of a bilayer with a pipette. Changes in transmembrane turgor or other mechanical pressures cause membrane tension. Yet even in the absence of these changes, Mscs can be activated by anesthetics, phospholipids missing one fatty acid (called lysophospholipids), and certain polyunsaturated lipids. These stimuli also perturb the membrane bilayers. Given the pore size, these proteins are less selective to ion flow compared to voltage-gate channels (see next section). Depending on the amino acids that line the pore, some Mscs would allow the preferential flow of anions while some allow cation flow. Some somatosensory channels (i.e. not pore) proteins also respond to pressure. When open, these can be selective to specific ions like Na+ or K+. Examples include some variants of the Transient Receptor Potential (TRP) ion channels. Other membrane proteins can also be activated by physical force. but true Mscs have some key characteristics. If mutated or deleted, the mechanosensory response is removed. If added to a cell, a mechanosensory response results. a. Small-conductance mechanosensitive channel This protein is a homoheptamer with three helices from each monomer involved in the overall structure. Two (helices 1 and 2) interact more with the lipid components of the bilayer. Interactions of specific lipids with the helices seem to promote closing but changes under high pressure lead to the opening of the pore. Half of each helix 3 forms the pore, while the other half is more parallel to the membrane and interacts with a large cytoplasmic domain. Figure $6$ shows the differences between the closed form of E. Coli MscS ( 2oau) and the open form (2vv5) viewing down the pore axis. The heptameric protein is shown in gray. Two key valines (105 and 109) on each chain are shown in spacefill and colored cyan. These hydrophobic amino acids act like gate-keepers, helping to keep water out, acting like a "vapor seal". In the closed state, the pore is sealed by the closing of the leucine "rings" as one half of helix 3 pack more closely. Many members of the MscS family vary significantly in size and can have between 3-11 transmembrane regions. The closed pore has a diameter of about 4.8 Å, while the open pore is 13 Å across. Most of you would have studied Ohm's law, given by \mathrm{V}=\mathrm{IR} \text { or } \mathrm{I}=\frac{\mathrm{V}}{\mathrm{R}} where I (amps) is the current, R (ohms) is the resistance and V (volts) is the voltage. A more general variant of this law used in physics is \mathrm{J}=\sigma \mathrm{E} where J is the current density, E is the electric field, and σ is the conductivity (inverse of resistance) which depends on the material. The unit of sigma σ is ohms-1 or mhos (ohm written backward). That unit has been renamed the Siemen (S). Mscs channels have a small conductance of approximately 1 nS in 400 mM salt solution. b. Large conductance mechanosensitive ion channel (MscL): The channels have large conductances (3 nS) and concomitantly larger pore sizes, allowing the flow of water, ions, and even small proteins. Again they are involved in diffusion down an electrochemical gradient through pores so not active transport just gated diffusion. Figure $7$ shows an interactive iCn3D model of the pentameric MscL from Mycobacterium tuberculosis (2oar), a Gram-positive bacteria that causes tuberculosis. About 23% of the world's population is affected by this pathogen. It causes about 1.5 million deaths each year. Compare this to the total number of deaths during the COVID pandemic (2020-21) of over 3 million (as of May 2021). It has killed over 1 billion people throughout human history (but not as many as malaria). The pore diameter is about 30 Å across. Pore-forming alpha-helical toxins We started this chapter section with a discussion of the major attack complex (MAC) of the innate immune system. Pathogens also employ pore formation to kill host cells. Many secrete soluble proteins which aggregate in the membrane to form either alpha-helical or beta-barrel pores. The proteins are called pore-forming toxins (PFTs). Killing occurs when either cytoplasmic components leak out or bacterial toxins, such as diphtheria and anthrax toxin, move into the cell. Figure $8$ shows an interactive iCn3D model of the pore formed by cytolysin A (ClyA, also known as HlyE), an alpha-PFT used by some E. Coi and Salmonella enterica strains. The pore is a large dodecamer that forms from soluble monomeric ClyA (2wcd). It has a pore diameter of about 40 Å, Figure $9$ shows the soluble monomeric form of cytolysin A (ClyA, HlyE) (1QOY). Nonpolar side chains are shown in cyan. The transparent surface is mostly polar, making the monomer soluble. Figure $10$ shows the changes in conformation between the single chain soluble form (shown in the figure above) and a single chain of the membrane oligomeric channel. We started this chapter section by exploring electroporation and the formation of a toroidal-like hole in the lipid bilayer. In the case of ClyA, the protein aggregate itself forms the pore, not the lipids themselves, although lipid rearrangements are necessary to form the protein complex. Gap Junctions Connexins are voltage-gated channels that allow for the flow of ions, metabolites, nucleotides, and small peptides. A connexin has four transmembrane helices and two extracellular loops, Six protomers of these come together in a single cell to form a channel complex called a connexon or hemichannel. Beta structures in the connexin hemichannel of one cell docks with a similar channel on an adjoining cell to form a full channel passing through the membranes of both cells forming a gap junction between the cells. This is shown in Figure $11$. There are 20 different connexins encoded in the human genome. The left figure below shows the six protomers, each in a different color and a gray rectangle representing the bilayer of a single connexon or hemichannel. The right side of the figure shows a full gap junction channel between two cells, with the membranes represented by gray rectangles. The connexin 26 monomer was used to create the diagram. Mutations in this protein are associated with hearing loss. The left figure above shows the six protomers, each in a different color and a gray rectangle representing the bilayer of a single connexon or hemichannel. The right side of the figure shows a full gap junction channel between two cells, with the membranes represented by gray rectangles. The connexin 26 monomer was used to create the diagram. Mutations in this protein are associated with hearing loss. The cytoplasmic entrance of the channel is positively charged. It forms a funnel, composed of 6 amino-terminal helices, which leads into a negatively charged transmembrane lining. The entrance diameter is 14 Å. Here is a model of a full gap junction channel connecting two membranes using the human connexin 26 monomer (2zw3). Only the bottom membrane bilayer is represented by red and blue dummy atoms. Figure $12$ shows an interactive iCn3D model of a full gap junction channel connecting two membranes using the human connexin 26 monomer (2zw3). Only the bottom membrane bilayer is represented by red and blue dummy atoms. The Nuclear Pore Complex (NPC) Channels have pores that can be gated open and allow the selective flow of ions. Pore-forming proteins have larger entrances that allow both small and large molecules to pass through the bilayer. The pore opening in even large mechanical sensitive channels (about pale in size compared to the nuclear pore complex, which has a combined molecular mass of around 125,000,000! Its outer diameter of ~1,200 Å and its inner one of about 425-Å. Figure $13$ shows the relative size of nuclear pore compared to other molecular structures including the eukaryotic ribosome, nucleosome, a soluble tetrameric protein (rubisco, 270K), and MscL (shown as a circle which represents the pore diameter). Its job is to shuttle small molecules by passive diffusion down a concentration gradient through the pore. In addition, it moves large molecules and molecular structures (proteins, RNA, and perhaps ribosomes) across the nuclear membrane in a process that requires energy. The proteins that comprise this complex are called nucleoporins (nups), of which there appear to be around 34 in humans. Each NPC complex contains around 1000 nucleoporins. The complex fuses the inner and outer nuclear membranes. We have focused so much on single bilayer membranes that comprise the plasma membrane and membranes of organelles like the Golgi complex and lysosomes, it might come as a surprise (perhaps not to biology students) that the nuclear membrane or envelope appears to consist of two bilayers. Most know that the mitochondria have two bilayers, an inner and outer membrane, similar to Gram-negative bacteria. Mitochondrial are believed to have arisen from bacteria so the double bilayer there makes sense. Figure $14$ shows the nuclear membrane or envelope of two bilayers (1) with an outer ring (2), spokes (3), a basket (4), and filaments (5). The NPC spans both bilayers. The outer bilayer of the nuclear envelope is continuous with the endoplasmic reticulum as shown in Figure $\PageIndex{15$ below. The dots on the ER membrane are ribosomes, making this the rough ER (as opposed to smooth ER, which has no attached ribosomes). Figure $\PageIndex{16$ below shows a model of the basket-like structure of the nuclear pore complex (NPC). It, as well as Figure XX, shows that instead of two separate bilayers, in actuality, there is just one bilayer with each leaflet bending around at the NPC and reversing directions! Think of the interesting lipids and protein components that enable the bend! Alternatively, you could say that 2 different membranes fuse at the NPC. The NPC consists of 32 copies of each specific nucleoporin (Nup) except two. One of those has 48 copies and the other 16 (even these sum to 2x32 Nups). Three rings form and surround the pore. There is a 16-membered ring of Nups facing the cytoplasm (cytoplasmic ring) and another 16-membered ring of Nups facing the nucleoplasm (nuclear ring). There is 8-fold rotational symmetry in each ring, suggesting a dimeric repeat of Nups in the rings. Eight Nups in the cytoplasmic ring have a disordered end that sticks out into the cytoplasm as filaments. In contrast, the disordered ends of eight Nups in the nuclear ring form filaments which bind together to form a ring at the bottom of the nuclear basket. The inner ring (called the FG Nups layer in the figure above), between the cytoplasmic and nucleoplasmic rings, also consists of Nups with ordered domains and disordered parts. The disordered parts of the inner ring Nups have repetitive sequences enriched in phenylalanine and glycine (hence the name FG Nups) and stick out into the central pore. These disordered region act as a filter allowing certain molecules to pass and excluding those greater with molecular weights greater than 40K. Large molecules need transport proteins called karyopherin transport factors to move through the pore. The core structure of the NPC, obtained through cryoelectron microscopy, is shown in Figure $\PageIndex{17$ below. The double bilayers are very evident. The nuclear and cytoplasmic filaments, as well as the disordered FG repetitive sequences that stick into the pore from the inner ring, are not seen since they are very flexible and don't' adopt single conformations using standard structure determination methods. Figure $18$ shows an interactive iCn3D model of the nuclear pore complex (NPC) core (5a9q), whose structure was obtained using cryoelectron microscopy. (load slowly) A cartoon figure showing the variety of Nups in the NPC is shown in Figure $19$. In this figure, the cytoplasmic and nucleoplasmic rings are both shown in green, each mostly formed by 16 copies of the Y-complex, arranged in two eight-membered rings. The inner ring, predominantly formed by 32 copies of the Nup93 complex is shown in red. Transmembrane nucleoporins are depicted in violet, and the cytoplasmic filaments and nuclear basket structure are in orange. Attached to the inner ring are Nup62 complexes (depicted in blue), which form a cohesive meshwork within the central channel through their FG-repeat domains. Not indicated is the position of Nup98, a FG-repeat-containing nucleoporin important for the transport and exclusion function of NPCs; its position in the NPC is less defined, but it might be part of the inner ring. Similarly, Aladin (also known as AAAS), Gle1, Rae1, and Npl1 (also known as hCG1 and NUPL2) have been omitted. Large proteins and RNA that pass through the pore must first be bound to a cargo receptor, which can move the "cargo" across the pore with concomitant GTP hydrolysis. This is a process that is closer to active transport so we will discuss that in Chapter 11.3. The entire nuclear pore complex was solved in 2022 using cryoEM. Here are two videos showing the dilated complex (7R5J). Click on the images to download mp4 animations of the complex.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/11%3A_Biological_Membranes_and_Transport/11.04%3A_Diffusion_Across_a_Membrane_-_Pores.txt

Search Fundamentals of Biochemistry In the previous sections, we explored facilitated diffusion, diffusion through channels, and diffusion through larger pores. In each case, once a carrier/permease protein was available, or a channel (gated by ligand binding, change in membrane potential, lipid binding, or mechanical forces) or a pore formed, solute flows down a chemical gradient (facilitated diffusion) or electrochemical gradient in a thermodynamically favored process. But what about moving solutes from low to high concentration, against a concentration gradient, which would be necessary to capture the last bit of a vital nutrient or energy source? Active transport does just that, but it requires an energy source to do so. Many different types of chemical species are actively transported across the cell and organelle membranes, including sugars, amino acids, (deoxy)nucleotides, metabolites (like carboxylic acids), Type of Active Transport For active transport to occur, a membrane receptor is required which recognizes the ligand to be transported. Of major interest to us, however, is the energy source used to drive transport against a concentration gradient. The biological world has adapted to use almost any source of energy available. ATP hydrolysis: One would expect that this ubiquitous carrier of free energy would be used to drive active transport. This is one of the predominant roles of ATP in the biological world. 70% of all ATP turnover in the brain is used for the creation and maintenance of a Na and K ion gradient across nerve cell membranes using the membrane protein Na+/K+ ATPase. Energy released by oxidation: You may have encountered this in previous biology courses. Active transport of protons driven by oxidative processes is exergonic. In electron transport in respiring mitochondria, NADH is oxidized as it passes electrons to a series of mobile electron carriers (ubiquinone, cytochrome C, and eventually dioxygen) using protein complexes in the inner membrane of the mitochondria. The energy lost in this thermodynamically favored process is coupled to conformational changes in the complex which caused protons to be ejected from the matrix into the inner membrane space. One can imagine a series of conformation-sensitive pKa changes in various side chains in the complexes which lead in concert to the vectorial discharge of protons. Light: Photosynthetic bacteria have a membrane protein called bacteriorhodopsin which contains retinal, a conjugated polyene derived from beta-carotene. It is analogous to the visual pigment protein rhodopsin in retinal cells. Absorption of light by retinal induces conformation changes in the retinal and bacteriorhodopsin, which leads to the vectorial discharge of protons. The collapse of an ion gradient: The favorable collapse of an ion gradient can be used to drive the transport of a different species against a concentration gradient. We have already observed that the collapse of a proton gradient across the inner mitochondria membrane (through F0F1ATPase) can drive the thermodynamically unfavored synthesis of ATP. The collapse of a proton gradient provides a proton-motive force that can drive the active transport of sugars. Likewise, a sodium-motive force can drive the active transport of metal ions. Since the energy to make the initial ion gradients usually comes from ATP hydrolysis, ATP indirectly powers the transport of the other species against a gradient. Active transport can be divided into two classes: • primary - driven by "primary" energy sources, which for us means ATP or photons • secondary - driven by coupled transport (also called cotransport) of solutes down a concentration gradient, which provides the thermodynamic force to move a solute "uphill". Active transporters can also be divided into classes based on the direction of movement of the solute and any cotransported solute into uniport (no cotransported solute), symport (solute and cosolute transported in the same direction) and antiport (solute and cosolute transported in the opposite direction) as shown in Figure \(1\). Examples of these different mechanisms can be found in facilitated diffusion as well as active transport. Examples of these types include: • uniport: An often-used example is GLUT 1, which catalyzes the FACILITATED DIFFUSION (not active transport) of glucose down a concentration gradient. A somewhat complicated example of a uniport active transporter is the Na+/K+ ATPase, which moves Na+ ions out of a cell and K+ into a cell, both against concentration gradients. The movement of one ion does not provide the chemical potential thermodynamic driving force to move the other. Rather the hydrolysis of ATP is required. So this protein can be called a uniport active transporter for two different ions. • symport: Another glucose transporter, the glucose symporter (SGLT1), found in the small intestines, heart, and brain, cotransports one glucose or galactose for every two Na+ ions that move into the cells down a concentration gradient. • antiport: The sodium-calcium exchanger pumps out one Ca2+ ion (low to high concentration) driven by the influx of three Na+ into the cell. This keeps Ca2+ low in the cytoplasm. If the species moved is charged, two other terms are used: • electrogenic - a net electrical imbalance is generated across the membrane by symport or antiport of charged species • electroneutral - no net electrical imbalance is generated across the membrane by symport or antiport of charged species In this chapter section, we will explore examples of several types of active transporters. All are polytopic with alpha-helical transmembrane domains, which through a series of conformational transitions can move chemical species that are bound to the receptor or channel protein to move through the membrane through the creation of transient openings in the protein. We will end with another look at the nuclear pore complex and see how it allows the movement of large proteins and RNA molecules through its pore through a very different type of mechanism. In that case, it is not active transport since the relative concentration of the transported species on either side of the pore is less important than the mechanism by which certain species are targeted to move into and out of the nucleus. Major Facilitator Superfamily (MFS) Transporters There are about 74 different families of active transporter in the Major Facilitator Superfamily (MFS), which is the largest of secondary transporters using symport or antiport mechanisms. There are also uniporters within this family. Different members of the MFS move a diverse array of solutes, including sugars, nucleotides, peptides, and drugs across membranes against a concentration gradient. All have two membrane domains consisting of six transmembrane helix bundles that pack to give an apparent two two-fold rotational axis of symmetry (also called pseudo C2 axis). The ligand to be transported binds between the two domains in a central cavity potentially open to either side of the membrane. Figure \(2\) panel A (top) below shows a cartoon of the 12 helices and 6-helix bundles. Helix 1 and 4 in domain one and the analogous ones in domains 7 and 10 are involved in ligand movement through the membrane. Correspondingly, helices 2, 5, 8, and 11 are involved in domain-domain interactions. The bottom figure (B) shows two views of the protein with the ligand in spacefill rendering. As structure mediates function, they all display a similar "clamp and switch" mechanism, in which solute binds to the protein in an open (to binding) conformation, leading to conformation changes forming a closed form, followed by further conformational changes which cause the complex to open to the other side of the membrane. Figure \(3\) shows hands opening, closing, and reopening in the other direction as an analogy for this "clamp-switch" mechanism. We will describe two different transporters in this family. Human monocarboxylate transporter (7BP3) These proton-linked monocarboxylate transporters (MCTs) move many monocarboxylates (including lactate, pyruvate, and the ketone bodies acetoacetate, beta-hydroxybutyrate and acetate) out of the cell, driven by the flow of protons across the membrane in the same directions (symport). In cells engaging in glycolysis, a fundamental energy utilization pathway, it catalyzes the high-affinity transport of pyruvate, the end product of glycolysis, and lactate, a reduced derivative of pyruvate, out of the cell. In cells that are not engaged in glycolysis, their activity is effectively turned off. The flux of pyruvate/lactate has steep dependence on their concentrations, allowing the protein to act as an on-offf switch for transport. A step dependency of activity on concentration is a hallmark of cooperative interactions. Structural analyzes show conformational changes in the interface of the dimeric form of the transporter. We will see when we study glycolysis, the conversion of the oxidized pyruvate to its reduced form lactate requires a reducing agent, in this case, NADH, which is oxidized back to NAD+, an oxidizing agent required for glycolysis to continue. When pyruvate is high, NADH is also high, allowing for the conversion of pyruvate to lactate, which can be exported out of the cell. The interconversion is described by this reaction: Pyruvate + NADH ↔ Lactate + NAD+ Using MCT 2 to remove lactate from the cell pulls the above reaction to the right, regenerating NAD+, and allowing glycolysis to continue. Figure \(4\) shows an interactive iCn3D model of the inward open human monocarboxylate transporter 2 (7BP3). Several key residues are shown. Aspartic acid 293 (side chains in red sticks) is conserved in all MCTs studied and is likely involved in H+ transport. It is surrounded by Val 156, Met 289, Ala 290, and Phe 351 (side chains in cyan sticks), all nonpolar amino acids which would elevate the pKa of Asp 293, allowing it to stay protonated until a conformational change would alter its environment, leading to proton release. Two positive charged residues, Lys 38 and Arg 297 (side chains in blue sticks) are involved in substrate interactions. Lys 38 faces to the outside of the cell in the inward open conformation. Figure \(\PageIndex{5\) below shows the same amino acids in a zoomed view. The amino acids are labeled. Lactose Permease (Lactose-proton symport) This E. Coli protein is also a symporter, which uses the movement of H+ down a concentration gradient to drive lactose into E. Coli to capture as much lactose - an energy source - as possible. Deprotonation of a protonated Glu 269 leads to ligand binding as the protein has no binding site in the protonated site. Lactose induces the formation of its binding site. Figure \(\PageIndex{6\) below shows the change in conformation going from a more acidic pH (5.6, PDB 2CFP) to a more neutral pH (6.5, PDB 2CFQ). One lactose is transported from one H+, which moves down a concentration gradient. Lactose moves from the outside periplasm to the cytoplasm. Figure \(7\) shows an interactive iCn3D model of lac permease (1pv7), which highlights three residues essential for lactose binding (Glu 126, Arg 144, and Glu 269 in stick) and three involved in proton transfer (Arg 302, His 322 and Glu 325 in spacefill). It has a bound nonhydrolyzable lactose analog (beta-D-galactopyranose-(1-1)-1-thio-beta-D-galactopyranose), shown in spacefill, in the center hydrophilic cavity. The two 6-helix bundles, which give pseudo 2-fold symmetry are evident. The protein is in the open-to-inward conformation. (The blue dummy atoms represent the inner leaflet.) Figure \(8\) is a video of a molecular dynamics simulation showing lactose moving through lactose permease. Figure \(8\): Molecular dynamics simulation showing lactose moving through lactose permease. https://www.ks.uiuc.edu/Gallery/Movi...annelProteins/ As with other members of the MFS transporters, lactose permease has 12 transmembrane helices that form two bundles. The lactose binding site is between the two bundles. When the protein is in the open-to-out conformation, it can bind protons, most likely at an exposed His. This enables lactose binding, which is followed by a conformational change to the open-to-in form. The protonated His loses a proton and lactose dissociates into the cytoplasm. Movement of both H+ and lactose together must occur for transport. Structural work by Singh et al on the Leu Transporter (LeuT), a member of the solute carrier 6 or sodium-coupled transporters, which is an active transporter requiring movement of Na ion into the cells to power the uptake of Leu, shows an "open-to-out" and occluded binding state for ligand (Leu). Tryptophan, a competitive nontransportable inhibitor binds to the open-to-out state but is too large for the obligate occluded state so it is not transported. Figure \(9\) shows inward open, closed, and outward conformations of different major facilitator superfamily transporters. Another member of the MFS Transporter family is LeuT, the leucine transport, which is similar to the neurotransmitter sodium symporter. Figure \(10\) shows the transition between two conformational states. The first is the outward open leucine transporter (LeuT). This initial state has bound tryptophan, a competitive inhibitor, which is too big to transport, which traps the bound state in the open conformation. The second state has bound Leu and is closed. The conformational changes are subtle but sufficient to allow transient binding, occlusion, and expulsion to the other side of the membrane (not shown). Note also the two bound Na+ ions (purple) clearly show that active transport through this transporter is driven by Na+ ions. The ions are farther apart in the open state allowing access to the amino acid for transport. Membrane ATPases Many active transporters power the uphill transport of solute through coupled exergonic cleavage of ATP. The largest family of is the P-type ATPase, which is used to transport a host of different solutes, including ions (Cu, Zn, Mn, Mg, Ca, Na, K, Cd, Co, Pb, Ni ,and H) and phospholipids (from one leaflet to another by translocases like flippases). They convert chemical energy (from cleavage of phosphoanhydride bonds) to electrochemical potential energy. Over 160,000 P-type ATPases from prokaryotes and eukaryotes are known so they are highly prevalent and evolved early in time. They should be understood at both a structural and thermodynamic level. How is energetic coupling achieved? An early model proposed by Albers and Post suggested that energetic coupling occurred by the covalent transfer of a terminal phosphate (Pi) from ATP to an Asp in the protein channel to form a covalent Asp-Pi intermediate, which is an example of a mixed anhydride. This is illustrated in Figure \(11\). This reaction is disfavored thermodynamically since hydrolysis of ATP to form ADP proceeds with a less negative ΔG0 than the same reaction with a mixed anhydride. However, ultimately hydrolysis of the mixed anhydride in the next step would make the overall coupled reactions favorable. Another feature of the Post-Albers model is that the phosphorylated channel, P-ATPase, has two conformations: • a high-affinity cation inward (cytoplasm) facing cation binding site called E1 • a low-affinity cation outward (extracellular or luminal) facing cation binding site called E2 Figure \(12\)s a more detailed (A) and simpler (B) set of chemical equations to represent the different states of E1 and E2 in the Post-Albers model. The next three figures are from the following reference: Zhang, X.C., Zhang, H. P-type ATPases use a domain-association mechanism to couple ATP hydrolysis to conformational change. Biophys Rep 5, 167–175 (2019). https://doi.org/10.1007/s41048-019-0087-1. Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/ • The top line in both shows the reactions of E1 (high affinity) with the substrate (ions) and ATP. Since E1 is phosphorylated by ATP, it is considered a kinase. • The bottom line in both shows the reactions of E2 (low affinity) with the substrate (ions). Since E2 is dephosphorylated, it is considered a phosphatase. S1 and S2 differ for specific P-type ATPase. Detailed Structures Now let's look at the structures of a specific P-type ATPase, the sarcoplasmic/endoplasmic reticulum calcium ATPase 1 (or more simply a Ca2+ pump), which catalyzes the active uphill transport of Ca2+ from the cytoplasm back into the lumen of the sarcoplasmic reticulum (SR) in muscle cells and acts in the regulation of striated muscle contraction. The actual free concentration of Ca2+ in the SR is about 390 μM compared to 0.1-0.2 μM in the cytoplasm. On muscle excitation, Ca2+ floods out into the cytoplasm but must be actively transported back into the SR by the Ca2+ pump. Here is the simplified chemical reaction: ATP + Ca2+(cyto) + H2O → ADP + Ca2+(s. rect) + H+ + Pi. We'll stick with a cartoon representation of each of the various structure states of this calcium pump, as illustrated in Figure \(13\) (also from Zhang et al). Again, different states of E1, the kinase state, are shown in the top line, while E2, the phosphatase state, is shown in the bottom line. PDB IDs are shown for each state. Ca2+ is shown as a cyan sphere. Note how the cytoplasmic N (nucleotide-binding), P (phosphorylation), and A (actuator) domains and the single transmembrane domain, colored to show the C-terminal (CTM) and N-terminal (NTM). are represented in the cartoon version to the right. P represents the phosphorylated Asp 351 mixed anhydride. The asterisk * denotes a transition state for the phosphorylation of Asp 351 (top line) and its dephosphorylation (bottom line). Ca2+ moves from the cytoplasm to the lumen of the SR. Note that the first E1 state is open inward (to the cytoplasm) while the first E2 state is open outward (to the lumen of the Sr, where the bulk of the Ca2+ ions are stored. This mimics the open-in and open-out states of the Major Facilitator Superfamily (MFS) channels discussed above. Detailed Thermodynamics Zhang et al provide two different views of the thermodynamics of Ca2+ ion pumping into the SR as shown in A (domain dissociation) and B (classic) in Figure \(14\). A quick view of the two "energy landscapes" show: • Both go from higher to lower free energy as expected since ATP hydrolysis powers the process. • They differ mostly in the energetic coupling steps In A, the actual energy released in ATP hydrolysis occurs when the Asp 351-Pi mixed anhydride is dephosphorylated (ΔGdeph), which is coupled to the dissociation of the P and A (* actuator) domains. This is shown in the structural cartoon near the last step in the complete cycle. In the cartoon diagram, 6 states are represented. Any mechanism can be broken down into more and more states on a more complete description of the actual mechanism. For example, ATP cleavage to ATP can be represented by three states, ATP, ADP, and Pi, or many more if intermediates and transition states are included, the complete Ca2+ transport cycle in the thermodynamics diagram is broken down into 12 states, represented by 8 horizontal solid lines _____ and 4 dashed ------ lines. Vertical lines with arrows show transitions between states. • Green arrows show changes in free energy due to the relative stability (chemical potential) of ATP. • Red arrows → show changes in free energy due to the relative stability (chemical potential) of the substrate ions • Cyan arrows show changes in free energy due to the electrical potential of the substrate ions ΔGL are for steps involving loading of reactants, while ΔGR are for release. In both diagrams, the conversion of state E1.S.ATP to E1P.S is uphill as we predicted for the formation of the On the far right of each graph are identical sets of large unfilled arrows (since ΔG is a state function) as the beginning and ending free energies of free E1 must be pathway (i.e. mechanism) independent. The two sets of upward unfilled arrows (red for the change in the chemical potential of the substrate ions and cyan for the change in the electric potential of the substrates are both positive as the ions move uphill from a lower to high electrochemical potential. The unfilled downward green arrow shows the change in free energy (or chemical potential) for ATP hydrolysis. Since energy cannot be created or destroyed, the narrow black arrow (Qx) represents energy that is dissipated in the reaction cycle. The starting and ending states are identical (i.e., E1), only being differed by the energy dissipation (denoted as QX) of the P-ATPase transporter during one functional cycle. The are other types of membrane ATPase whose structure and function differ from the P-type described above. Some run in reverse to synthesize ATP. They also vary in substrate ions. Here are several different types. • F-ATPases (ATP synthases, F1F0-ATPases): These are used to synthesize ATP and are powered by the collapse of a H+ gradient. They are found n mitochondria, chloroplasts, and bacterial cell membranes. We will discuss these more in chapters on mitochondrial oxidative phosphorylation and photosynthesis in chloroplasts. • V-ATPases (V1V0-ATPases): These are used to pump protons into organelles to acidify them (ex. lysosome) and are also found in bacteria. • A-ATPases (A1A0-ATPases): These are used to synthesize ATP in Archaea. • E-ATPases: These are found on the cell surface and hydrolyze extracellular nucleotide triphosphate. Let's now look at some special features of one P-type ATPase that transports both Na+ and K+ ions and is important in establishing their intracellular and extracellular concentration in neurons, so they are critical for neuron function. Na+/K+ ATPase This protein keeps the K+in and Na+out high compared to their respective concentrations on the other side of the neural cell membranes. ATP and 3 Na+ ions bind to the cytoplasmic domain of the enzyme in the E1 conformation. As described more generally above, in the presence of Na ions, the bound ATP is cleaved in a nucleophilic attack by an Asp side chain of the protein. Hence, the protein is a Na+-activated ATPase or kinase. The phosphorylated enzyme changes conformation to the E2 form in which Na+ ions are now on the outside of the cell membrane, from which they dissociate. The phosphorylated protein in conformation E2 now binds 2 K+ ions on the outside, which activates hydrolysis of the Asp-PO3 mixed anhydride link. The dephosphorylated protein is more stable in the E1 conformation, to which it changes as it brings K+ ions into the cell. Hence this protein is an electrogenic antiporter. P-Type transporters are inhibited by vanadate (VO43-), a transition state analog of phosphate. Transport mediated by P-type membrane proteins can, in the lab, be used to drive ATP synthesis. Detailed kinetic analysis of ATP and VO43- interactions show there are low-affinity and high-affinity sites for each on Na/K ATPase. The high-affinity vanadate site appears to be the same as the low-affinity ATP site, which suggests that vanadate binds tightly to the E2 form of the enzyme. The low-affinity vanadate site appears to be the same (based on competition assays) as the ATP site, which is probably the E1 form. Hence vanadate binds preferentially to the E2 form which would inhibit the transition to the E1 form. Vanadate also inhibits phosphatases, enzymes that cleave phosphorylated Ser, Thr, and Tyr phosphoesters in proteins. This supports the notion that vanadate binds preferentially to the E2 form, which has a phosphoanhydride link (Asp-O-phosphate) that is hydrolyzed, promoting the conversion of E2 back to E1. Vanadate is probably a transition state analog inhibitor in that it can readily adopt a trigonal bipyramidal structure, mimicking the transition state for cleavage of the tetrahedral anhydride bonds of ATP and Asp-O-PO3. Figure \(15\) is a YouTube animation of the Na+/K+ ATPase from xx. Permission Question? Figure \(15\): Animation of the Na+/K+ ATPase pump These interactions are depicted in Figure \(16\)below (after Stryer 4th ed) ABC Transporters The proteins comprise one of the largest families of membrane proteins with seven main families (ABCA to ABCG). Up to 3% of all bacterial proteins encode proteins associated with the ABC transporters. Different ABC transporters move a variety of required chemical species, from small (ions, sugars, amino acids, nucleosides, vitamins) to large (peptides, lipids, oligonucleotides, and polysaccharides) into the cell. They also remove toxic (to the cell) molecules such as xenobiotics (molecules foreign to the cell like drugs, toxins, etc) and potentially toxic metabolites. All of these require ATP hydrolysis. Moving toxic molecules out of the cell is beneficial to the health of the cell, but in the case of a tumor cell, not to the benefit of the organism. TAs in other active transporters using ATP, its hydrolysis leads to two different conformations, open-outward and open-inward. All ABC transporters have a LSGGQ amino acid sequence in the NBD. They also have a phosphate-binding loop (P-loop or Walker A motif). Most eukaryotic ABC transporters move solutes from the inside to the outside of the cell. The various structures within the ABC transporter superfamily are shown in Figure \(17\). The transmembrane domain (TMD, in green) and the nucleotide-binding domain (NBD, blue) are present in most versions of the gene. PK represents prokaryotic and EK eukaryotic organisms (shown at the very right for each). The ABC transporter genes denoted as full structures have 2 TMDs and 2 NMDs while half structures have one of each. Some of the genes encode either a single NBD or a single TMD (prevalent in prokaryotes, along with half structures). The ABC2 structure has only two NBDs. The single structure represents the ABC transporter gene with a single TMD or NBD; ABC2 structure represents the ABC transporter gene with only two NBDs. The families possessing certain structures are listed on the right. TMDs typically have 6-10 transmembrane α-helices. Those that are involved in the export of chemical species have six. Structural cartoons representing domain and ligand binding for a variety of ABC transporters are shown in Figure \(18\) below. (A) shows an inward-open drug (D) exporter. Binding of 2 ATPs cause dimer formation between the two NBD domains, resulting in a conformation change to outward facing which allows dissociation of the drug. Hydrolysis of ATP enables the release of ADP/Pi and dissociation of the 2 NBD domain, which results in conformational change to the inward-open form. (B) shows the delivery of a substrate (S) from its binding protein (ex. part of the ABC transporter complex MalEFGK involved in maltose/maltodextrin import) in the periplasm of E. Coli for delivery into the cell. (C) shows an outward-open variant of (B) (ex. part of the Vitamin B12 import system permease protein BtuC) The mammalian protein multi-drug resistance (MDR), also known as P-glycoprotein, Phospholipid transporter ABCB1 or ATP-dependent translocase ABCB1, is an example of an ABC Transporter. It moves both drugs across the membrane as well as phospholipids, including phosphatidylcholine, phosphatidylethanolamine, ceramides, and sphingomyelins, from the inner to the outer leaflet of the membrane. The gene for this protein is often mutated in tumor cells which moves chemotherapeutic drugs from the cell. Figure \(19\) shows an interactive iCn3D model of the structure of the mouse P-glycoprotein (4M2T) bound to a cyclic peptide inhibitor in the inward open conformation is shown below. Most proteins bind substrates specifically but P-glycoprotein binds them quite indiscriminately which makes this protein so useful in pumping drugs out of the cell. The iCn3D model below shows the conserved (mouse and human) transmembrane domain aromatic amino acids (H60, F71, T114, F299, Y303, Y306, F332, F339, F724, F728, F766, F938, Y949, F953, F974, F979) involved in the transport pathway in colored sticks. The inhibitor (cyclic-tris-(S)-valineselenazole; QZ59-SSS) is shown in spaceiflll. The consensus LSGGQ (527-531 and 1172-1176) sequences in the nucleotide-binding domain are shown in colored spheres. Here is a view showing just the transmembrane domain with the conserved aromatic amino acids again. Figure \(20\) shows an interactive iCn3D model of just the transmembrane domain with the conserved aromatic amino acids of the mouse P-glycoprotein bound to a cyclic peptide inhibitor in the inward open conformation (4M2T). Another look at the Nuclear Pore Complex We examined the incredibly complicated structure of the nuclear pore in the previous section. Both small molecules and large proteins (synthesized in the cytoplasm) and RNAs (synthesized in the nucleus) move through its large pore. The movement of solutes through its gaping pore does not require either ATP cleavage in a primary active transport or the collapse of a chemical gradient in a secondary active transport process. Then why study it in this chapter section? It turns out that the regulation of the process requires GTP cleavage and a gradient of a particular protein called Ran. What's fascinating about the nuclear pore is its selectivity for protein transfer across the pore. What proteins are allowed in and out? What is the origin of the specificity? The specificity is determined in part by a protein family called the karyopherin-βs (22) of receptors, which bind and transport nuclear proteins. There are two types of these nuclear transport receptors or transport factors: • Importins facilitate the movement of proteins into the nucleus (ex. nuclear proteins like histones, DNA and RNA polymerases, etc). • Exportins facilitate the movement of RNA (except mRNA) out of the nucleus. Large proteins destined to be moved through the nuclear pore are called cargo proteins. They have a molecular signal that differentiates them from molecules that should stay in the cytoplasm or move into organelles or be secreted from the cell. The signals are called: • NLS or nuclear localization sequences • NES or nuclear export sequences There is no obvious NLS consequence and different motifs are used by different cargos. A classical NLS is enriched in Lys and Arg residue. Others appear enriched in Pro-Tyr or Ile-Lys. Large domain structures might also participate in the NLSs. Prediction programs are used to identify clusters of lysines and arginines with gaps between the clusters. Classical NESs appear to be rich in leucines in an 8-15 amino acid sequence with regularly spaced conserved hydrophobic amino acids. A third player, RAN (Ras-related nuclear protein), is involved that determines which way the transport receptor:cargo complex moves. Ran is a small protein that binds and can hydrolyze GTP to GDP. It is a member of the small G proteins that are GTPase. Ran itself can run (a pun) or move across the nuclear membrane and is found in both the cytoplasm and nucleus. What determines which way a Cargo:receptor complex goes? It depends on whether GTP or GDP is bound to RAN! There are higher concentrations of RanGTP in the nucleus and lower concentrations in the cytoplasm. • Importins bind cargo proteins with a NLS in the cytoplasm where RanGTP is low, move into the nucleus, and release the cargo protein when the abundant RanGTP displaces the bound cargo protein • Exportins bind both a cargo protein with a NES and Ran:GTP to form a ternary Exportin:Cargo:RanGTP complex in the nucleus. This moves into the cytoplasm, where the Ran bound GTP is hydrolyzed to GDP, causing the complex to dissociate, freeing the exported cargo protein from the complex. Importins are dimeric structures consisting of an alpha and beta subunit. The alpha subunit binds to the cargo protein through the NLS sequence on the cargo protein. The beta subunit interacts with the nucleoporin proteins (Nups) in the These interactions are shown in Figure \(21\). Figure \(22\) shows an interactive iCn3D model of the Delpi electrostatic surface potential map of the putative NLS from the carboxy-terminal of a cargo protein, the W protein of the Hendra virus (4M2T)which is imported into the nucleus. The virus derives from bats and has recently emerged. The peptide sequence containing the NLS has the following sequence: 419 CLGRRVVQPGMFADYPPTKKARVLLR 444. The red surface indicates the negative potential and the blue positive potential, which is associated with the Lys and Arg side chains in the consensus sequence. It is shown bound to the human importin-α3 subunit (molecular surface shown in white), which binds the NLS sequence with high affinity. An astute reader might ask why GTP stays bound and is not cleaved into GDP in the nucleus by the intrinsic GTPase activity of Ran. It turns out there is yet another protein found only in the cytoplasm (i.e. it doesn't have a nuclear import signal), which binds to Ran:GTP and promotes GTP → GDP exchange. So oddly, hydrolysis of a nucleotide triphosphate (GTP, not ATP as in the case of most transporters discussed above) is required for nuclear import. A different but very fascinating mechanism! The cargo complexes described above must still pass through the nuclear pore. In section 11.3 we discussed the structure of the nuclear pore which consists of many nucleoporin proteins (NUPs). The inner ring is called the FG Nups layer. The disordered parts of the inner ring Nups have repetitive sequences enriched in phenylalanine and glycine (hence the name FG Nups) and stick out into the central pore. The FG repeats can take multiple forms including Phe-Gly (FG), Gly-Leu-Phe-Gly (GLFG), or Phe-any-Phe-Gly (FXFG). These are intrinsically disordered proteins and the disordered regions in the pore act as a filter allowing certain molecules to pass and excluding those greater with molecular weights greater than 40K. Large molecules complexed to importins/exportins move through the pore. This disordered mesh prevents passive diffusion of molecules through it but allows protein complexes with cargo:exportin/importin complex through. It's a bit like electrophoresis of small protein complexes through the pores of an acrylamide or agarose gel polymerized matrix, only without the "push" of an electric field. Presumably, the FG-Nups make transient hydrophobic (induced-dipole:induced dipole) interactions between the FGs on the Nups and the nuclear transport receptors. Figure \(23\) shows an export complex (5XOJ) from yeast of Ran (cyan with bound GTP in spacefill) bound to an exportin (Xpo1p, white surface) and 3 Nup42p peptides containing SxFG/PxFG repeats (spacefill, side chains, which are mostly nonpolar). To bind 3 Nup peptides, the exportin Xpo1 also contains repeating binding sites on domains called HEAT repeats 14–20 of Xpo1p, The exportin Xpo1p is shaped like a toroid with 21 HEAT repeat domains that have two antiparallel sheets with connecting loops of different sizes. Some believe the unfolded mesh of FGs in the core might condense on the binding of motifs on importins/exportins or through FG:FG domain associations. But what is the basis for importins/exportins interactions with the FG structures? To study it in more detail Fragasso et al designed and made an artificial FG-Nup that binds to an importin transport receptor Kap95 that interacts with a cargo protein with a nuclear localization sequence (NLS) in cargo proteins. They attached them to the inside of solid-state nanopores and demonstrated the fast movement of Kap95 through the derivatized nanopore while a control "non cargo" without a NLS, BSA) was blocked. Underivatized nanometer-sized pores are made in a silicon nitride synthetic membrane (SiN) by using an ion or electron beam to tune the size of the hole. Figure \(24\) (top image) shows colored code structures of the inner ring (top of figure) snapshots of different yeast GLFG NUPs. These are named after their Gly-Leu-Phe-Gly repeating motifs and are especially cohesive. The GLFG-Nups, shown in red, are mostly found in the inner ring compared to FxFG/FG-Nups shown in green. The bottom part of Figure \(24\) shows snapshots of molecular dynamics simulations of various yeast GLFG-Nups compared to the NupX synthetic one. Three of the yeast GLFG Nups and the synthetic NupX show a compact and extended domain Each has a collapsed cohesive domain at one end (characterized by a low charge to hydrophobic amino acid residue ratio (C/H ), lots of alternating FG and GLFG repeats, with light green showing non-FG/GLFG/charged residues ) and an extended domain at the other end (high ratio of charged/hydrophobic amino acids, no FG repeats, pink-red showing non-FG/GLFG/charged residues). Bright green shows the FG repeats, red the GLFG repeats, and white the charged groups. Figure \(25\) shows the pore in the silicon nitride synthetic membrane (SiN) membrane before (left) and after derivitization with NupX. Figure \(26\) shows derivatized pores in the SiN membrane of increasing size. In the smallest particle, the NupXs don't fit and are expelled from the pore. In the 30 nm particle, the NupX cohesive domains plug the hole. In the 45 nm pore, a hole appears. Figure \(26\): Derivatized pores in the silicon nitride (SiN) membrane of increasing size Lastly, Figure \(27\) shows snapshots of 30 nm pores lines with NupXs. In panel c, an importin, Kap95 (spheres with orange binding spots) is shown forming transient interaction with and translocating through the NupX-lined hole. BSA (sphere without binding sites) may interact weakly but does not translocate through the pore.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/11%3A_Biological_Membranes_and_Transport/11.05%3A__Active_Transport.txt

Search Fundamentals of Biochemistry Pores and Pore-Forming Proteins (PFPs) If you form a pore in a cell bilayer, molecules of all sizes could move either way based on their electrochemical potential. They will move from regions of a higher to lower electrochemical potential in a thermodynamically favorable process. Hence movement through pores represents a special case of facilitated diffusion when part of the driving force is not just a concentration gradient but also an electrical potential. Several questions might come to mind. • What proteins are involved in pore formation? • How is the specificity of solute movement through the pore regulated? • What is the mechanism of pore formation? Pore formation can lead to cell death, which is the function of some pore-forming proteins (PFPs) including the toxin Hemolysin E (also known as HlyE, ClyA, SheA) secreted from E. Coli and S. Aureus. Human proteins also form a membrane attack complex (examples include the membrane attack complex-perforin/cholesterol-dependent cytolysin (MACPF/CDC) superfamily and the membrane attack complex (MAC). The MAC consists of an assembly of proteins involved in the complement system (part of the effector branch in the innate immune system), which leads to cell death of Gram-negative bacteria like E. Coli. Figure $1$ illustrates the assembly process of the membrane attack complex and the complexity of interactions required to form a lethal pore in a cell. Complement protein C9 can adopt a soluble form or membrane form which in aggregates form the pore leading to cell death. This is a common feature of PFPs. PFPs could create a pore by altering membrane lipid packing to form a toroid-like hole (Figure $2$) and/or by inserting in a membrane and forming a pore within the protein complex itself. In either mechanism, lipid packing is altered. Biophysical evidence shows some support for the formation of "toroidal pores". Lipid rearrangements in the membrane could lead to a hydrophilic or hydrophobic pore lining as shown in Figure $3$. We started our study of lipid bilayers with pure lipid systems and then added membrane proteins. Let's do the same with pore formation. A common technique to form a pore in pure lipid bilayers and also in cells is electroporation. This is a technique used to move a DNA with a target gene into either a prokaryotic cell (transformation) or eukaryotic cell (transfection) for exogenous gene expression. In the absence of PFPs, this requires the alteration of surface tension by applying an electrical potential. This forms depressions in the membrane, altering the nonpolar acyl chain packing. In the process, small wire-like columns of water appear, which like hydrophobic pores ultimately rearrange into hydrophilic toroidal pores. Figure $4$ shows snapshots of molecular dynamics simulations as a function of the time of pore formation in electroporation. How does DNA pass through the pores in the bilayer? In pure lipid vesicles, it appears to pass through by electrophoresis. Most students are familiar with the electrophoresis of DNA fragments through pores in agarose gels. In actual living cells, small nucleic acids like small interfering RNA (siRNA) and antisense DNA molecules appear to pass through the bilayer by electrophoresis. Large DNAs like plasmids containing a gene for expression bind to the cell and form cell surface aggregates, which appear to be endocytosed into the cell. Electroosmosis, the movement of liquids under the influence of an electric field, also plays a role. Pores - Outer Membrane Factor (OMF) and Voltage-Dependent Anion Channel (VDAC) Now let's consider pores made of PFPs. We have already discussed two types of beta-barrel transmembrane proteins, the outer membrane factor (OMF) of Gram-negative bacteria and the voltage-dependent anion channel (VDAC). Both are examples of proteins called porins with typical beta-barrel topology. VDAC: At low membrane potentials, VDAC (also known as mitochondrial porin), the most abundant protein in the mitochondrial outer membrane, moves metabolites and Ca2+ ions across the outer membrane of mitochondria. VDAC exist in an open state at 0 or very low transmembrane potentials that allows for the transfer of key metabolic anions (pyruvate, oxaloacetate, malate, succinate, ATP, ADP, and Pi, which are involved in metabolism) and Ca2+, and in a closed state (above or below + 30 mV), which is not completely closed as it allows for the transfer of ions with a preference for cations. The closed state does not allow for the transfer of ATP. The transition to the closed state is promoted by tubulin and actin (cytoskeletal proteins), negatively charged lipids such as phosphatidyl ethanolamine and cardiolipin, and also covalent phosphorylation by protein kinases. Bcl2, proteins involved in the regulation of programmed cell death (apoptosis) also interact with VDAC. In contrast to ligand-gated channels which require ligand binding to open the channel, pore complexes are unusually open. Millions of ATPs/second move across the membrane through the open pore but none through the closed pore. Figure $5$ shows an interactive iCn3D model of mouse VDAC1 (4c69) with bound ATP loosely held in the site. An alpha helix partly occludes the central pore of this β-barrel protein. One ATP is bound in the barrel and interacts with Lys 12 and Lys 20 at each end of the cavity-bound helix. The alpha-helix narrows the pore opening and presumably changes orientation in a voltage-sensitive fashion to gate the pore open and closed, hence regulating the conductance of ions through the pore. That the orientation of the charged arginine side chains would be dependent on the transmembrane potential should be somewhat obvious. Aquaporins Aquaporins can move a billion water molecules per second across membranes and exclude ions including protons. Waters proceed in a single file through the pore. Instead of moving waters through, it might simply move "H+ ions" through by the alignment of subsequent H bond donors and acceptors in a "wire" of water molecules. This is prevented by two conserved asparagine residues in the center of the channel, which disrupt water to water hydrogen bond network in the channel waters that could facilitate proton transfer. Instead, the central waters form hydrogen bonds to the central asparagines. This, along with local membrane potentials, causes opposite orientations of the water in different leaflet sides of the membrane, precluding H+ transfer. Here is a movie of a molecular dynamics simulation of water moving through a porin called aquaporin, GlpF, from E. Coli. Science magazine (Tajkhorshid et al., Science Apr 2002, 296:525). Used with permission from the Theoretical and Computational Biophysics Group, the National Institutes of Health (NIH) Resource for Macromolecular Modeling and Bioinformatics, at the Beckman Institute, University of Illinois at Urbana-Champaign. OMF: The outer membrane factor (see previous section) is one member of a class of bacterial porins, which are the most abundant proteins in the outer membrane of Gram-negative bacteria. They are classified as non-specific or specific (with respect to the solute that passes through), or monomeric, dimeric, or trimeric based on their structure. In Gram-negative bacteria, which have two lipid bilayers, the movement of solute from inside to outside includes at least three sets of proteins. Active transport (discussed in the next section) needs an energy source and is used by inner membrane transport proteins, including ATP-binding cassette (ABC)-type, resistance nodulation division (RND)-type and major facilitator superfamily (MFS)-type transporters. These are connected to membrane fusion proteins (MFP) that span the periplasm, which then interacts with at least 21 different types of porins. Molecules with molecular weights greater than 600 generally can not get through the nuclear envelope of Gram-negative bacteria, limiting the size of potential antibiotics, which must enter by passive diffusion. Mechanosensitive ion channels - Mscs (which are pores!) As the name applies, these ion channels (with openings large enough to be called pores) are gated open/closed by mechanical (physical) changes in the properties of the membrane, not extracellular/intracellular ligands or voltage changes. Certain bilayer lipids also activate the Mscs. There are two types, small (MscS) and large (MscL) mechanosensitive ion channels. Changes in local (boundary layer) and nonlocal lipids are involved in the gating of the channel (in the next section we will discuss lipid-gated ion channels). They are found in prokaryotes, archaea ,and eukaryotes. They are also called stretch-gated ion channels. Mcss transduces a physical force (stretching and change in turgor pressure) into an electric signal - a flow of ions across the membrane. Turgor pressure is the internal pressure that "presses" the cytoplasm and cell membrane towards the cell wall in bacteria and plant cells. It arises mostly from the osmotic flow of water into the cell. If bacterial cells are placed in a high salt concentration solution (hypertonic), water flows out of the cell and the cell membrane shrinks to the inside of the volume confined by the cell wall. When placed in a hypotonic solution, water flows into the cell and the cell membrane presses out to the cell wall. The response of these channels is fast, in the millisecond range, which is about as quick as a cellular response can be. Some variants of these channels are called piezochannels, based on the piezoelectric effect that describes how a voltage is produced when some materials are deformed by mechanical stress which causes a redistribution of charges. Bacteria normally have high concentrations of both K+ and negatively charged anions, especially glutamate, which leads to a high turgor pressure from the inward osmotic flow of water. At low external osmolarity, turgor pressure in the cell could be as high as 4 atm. If placed in external solutions of high osmolarity, bacterial cells respond by the increased movement of solutes into the cell. Mscs are particularly important when bacteria are subjected to sudden osmotic shock. If they are placed in pure water, for example, water would flow down a concentration gradient into the cell and cause the cell to swell and lyse, killing the cell. This is done in the lab to prepare almost pure hemoglobin from ruptured red blood cells. The Mscs are opened under these conditions and small species from the cytoplasm flow out, helping to keep the cell viable. Their openings must be regulated to prevent too much outward flow, which would kill the cell. In other organisms they are also involved in touch (stretch), hearing (vibration, sound waves), and responses to gravity. Stimuli to activate them include fluid shear stress (relevant to endothelial cells that line blood vessels), membrane stretch (relevant to skeletal and cardiac muscle cells), or even indentation of a bilayer with a pipette. Changes in transmembrane turgor or other mechanical pressures cause membrane tension. Yet even in the absence of these changes, Mscs can be activated by anesthetics, phospholipids missing one fatty acid (called lysophospholipids), and certain polyunsaturated lipids. These stimuli also perturb the membrane bilayers. Given the pore size, these proteins are less selective to ion flow compared to voltage-gate channels (see next section). Depending on the amino acids that line the pore, some Mscs would allow the preferential flow of anions while some allow cation flow. Some somatosensory channels (i.e. not pore) proteins also respond to pressure. When open, these can be selective to specific ions like Na+ or K+. Examples include some variants of the Transient Receptor Potential (TRP) ion channels. Other membrane proteins can also be activated by physical force. but true Mscs have some key characteristics. If mutated or deleted, the mechanosensory response is removed. If added to a cell, a mechanosensory response results. a. Small-conductance mechanosensitive channel This protein is a homoheptamer with three helices from each monomer involved in the overall structure. Two (helices 1 and 2) interact more with the lipid components of the bilayer. Interactions of specific lipids with the helices seem to promote closing but changes under high pressure lead to the opening of the pore. Half of each helix 3 forms the pore, while the other half is more parallel to the membrane and interacts with a large cytoplasmic domain. Figure $6$ shows the differences between the closed form of E. Coli MscS ( 2oau) and the open form (2vv5) viewing down the pore axis. The heptameric protein is shown in gray. Two key valines (105 and 109) on each chain are shown in spacefill and colored cyan. These hydrophobic amino acids act like gate-keepers, helping to keep water out, acting like a "vapor seal". In the closed state, the pore is sealed by the closing of the leucine "rings" as one half of helix 3 pack more closely. Many members of the MscS family vary significantly in size and can have between 3-11 transmembrane regions. The closed pore has a diameter of about 4.8 Å, while the open pore is 13 Å across. Most of you would have studied Ohm's law, given by \mathrm{V}=\mathrm{IR} \text { or } \mathrm{I}=\frac{\mathrm{V}}{\mathrm{R}} where I (amps) is the current, R (ohms) is the resistance and V (volts) is the voltage. A more general variant of this law used in physics is \mathrm{J}=\sigma \mathrm{E} where J is the current density, E is the electric field, and σ is the conductivity (inverse of resistance) which depends on the material. The unit of sigma σ is ohms-1 or mhos (ohm written backward). That unit has been renamed the Siemen (S). Mscs channels have a small conductance of approximately 1 nS in 400 mM salt solution. b. Large conductance mechanosensitive ion channel (MscL): The channels have large conductances (3 nS) and concomitantly larger pore sizes, allowing the flow of water, ions, and even small proteins. Again they are involved in diffusion down an electrochemical gradient through pores so not active transport just gated diffusion. Figure $7$ shows an interactive iCn3D model of the pentameric MscL from Mycobacterium tuberculosis (2oar), a Gram-positive bacteria that causes tuberculosis. About 23% of the world's population is affected by this pathogen. It causes about 1.5 million deaths each year. Compare this to the total number of deaths during the COVID pandemic (2020-21) of over 3 million (as of May 2021). It has killed over 1 billion people throughout human history (but not as many as malaria). The pore diameter is about 30 Å across. Pore-forming alpha-helical toxins We started this chapter section with a discussion of the major attack complex (MAC) of the innate immune system. Pathogens also employ pore formation to kill host cells. Many secrete soluble proteins which aggregate in the membrane to form either alpha-helical or beta-barrel pores. The proteins are called pore-forming toxins (PFTs). Killing occurs when either cytoplasmic components leak out or bacterial toxins, such as diphtheria and anthrax toxin, move into the cell. Figure $8$ shows an interactive iCn3D model of the pore formed by cytolysin A (ClyA, also known as HlyE), an alpha-PFT used by some E. Coi and Salmonella enterica strains. The pore is a large dodecamer that forms from soluble monomeric ClyA (2wcd). It has a pore diameter of about 40 Å, Figure $9$ shows the soluble monomeric form of cytolysin A (ClyA, HlyE) (1QOY). Nonpolar side chains are shown in cyan. The transparent surface is mostly polar, making the monomer soluble. Figure $10$ shows the changes in conformation between the single chain soluble form (shown in the figure above) and a single chain of the membrane oligomeric channel. We started this chapter section by exploring electroporation and the formation of a toroidal-like hole in the lipid bilayer. In the case of ClyA, the protein aggregate itself forms the pore, not the lipids themselves, although lipid rearrangements are necessary to form the protein complex. Gap Junctions Connexins are voltage-gated channels that allow for the flow of ions, metabolites, nucleotides, and small peptides. A connexin has four transmembrane helices and two extracellular loops, Six protomers of these come together in a single cell to form a channel complex called a connexon or hemichannel. Beta structures in the connexin hemichannel of one cell docks with a similar channel on an adjoining cell to form a full channel passing through the membranes of both cells forming a gap junction between the cells. This is shown in Figure $11$. There are 20 different connexins encoded in the human genome. The left figure below shows the six protomers, each in a different color and a gray rectangle representing the bilayer of a single connexon or hemichannel. The right side of the figure shows a full gap junction channel between two cells, with the membranes represented by gray rectangles. The connexin 26 monomer was used to create the diagram. Mutations in this protein are associated with hearing loss. The left figure above shows the six protomers, each in a different color and a gray rectangle representing the bilayer of a single connexon or hemichannel. The right side of the figure shows a full gap junction channel between two cells, with the membranes represented by gray rectangles. The connexin 26 monomer was used to create the diagram. Mutations in this protein are associated with hearing loss. The cytoplasmic entrance of the channel is positively charged. It forms a funnel, composed of 6 amino-terminal helices, which leads into a negatively charged transmembrane lining. The entrance diameter is 14 Å. Here is a model of a full gap junction channel connecting two membranes using the human connexin 26 monomer (2zw3). Only the bottom membrane bilayer is represented by red and blue dummy atoms. Figure $12$ shows an interactive iCn3D model of a full gap junction channel connecting two membranes using the human connexin 26 monomer (2zw3). Only the bottom membrane bilayer is represented by red and blue dummy atoms. The Nuclear Pore Complex (NPC) Channels have pores that can be gated open and allow the selective flow of ions. Pore-forming proteins have larger entrances that allow both small and large molecules to pass through the bilayer. The pore opening in even large mechanical sensitive channels (about pale in size compared to the nuclear pore complex, which has a combined molecular mass of around 125,000,000! Its outer diameter of ~1,200 Å and its inner one of about 425-Å. Figure $13$ shows the relative size of nuclear pore compared to other molecular structures including the eukaryotic ribosome, nucleosome, a soluble tetrameric protein (rubisco, 270K), and MscL (shown as a circle which represents the pore diameter). Its job is to shuttle small molecules by passive diffusion down a concentration gradient through the pore. In addition, it moves large molecules and molecular structures (proteins, RNA, and perhaps ribosomes) across the nuclear membrane in a process that requires energy. The proteins that comprise this complex are called nucleoporins (nups), of which there appear to be around 34 in humans. Each NPC complex contains around 1000 nucleoporins. The complex fuses the inner and outer nuclear membranes. We have focused so much on single bilayer membranes that comprise the plasma membrane and membranes of organelles like the Golgi complex and lysosomes, it might come as a surprise (perhaps not to biology students) that the nuclear membrane or envelope appears to consist of two bilayers. Most know that the mitochondria have two bilayers, an inner and outer membrane, similar to Gram-negative bacteria. Mitochondrial are believed to have arisen from bacteria so the double bilayer there makes sense. Figure $14$ shows the nuclear membrane or envelope of two bilayers (1) with an outer ring (2), spokes (3), a basket (4), and filaments (5). The NPC spans both bilayers. The outer bilayer of the nuclear envelope is continuous with the endoplasmic reticulum as shown in Figure $\PageIndex{15$ below. The dots on the ER membrane are ribosomes, making this the rough ER (as opposed to smooth ER, which has no attached ribosomes). Figure $\PageIndex{16$ below shows a model of the basket-like structure of the nuclear pore complex (NPC). It, as well as Figure XX, shows that instead of two separate bilayers, in actuality, there is just one bilayer with each leaflet bending around at the NPC and reversing directions! Think of the interesting lipids and protein components that enable the bend! Alternatively, you could say that 2 different membranes fuse at the NPC. The NPC consists of 32 copies of each specific nucleoporin (Nup) except two. One of those has 48 copies and the other 16 (even these sum to 2x32 Nups). Three rings form and surround the pore. There is a 16-membered ring of Nups facing the cytoplasm (cytoplasmic ring) and another 16-membered ring of Nups facing the nucleoplasm (nuclear ring). There is 8-fold rotational symmetry in each ring, suggesting a dimeric repeat of Nups in the rings. Eight Nups in the cytoplasmic ring have a disordered end that sticks out into the cytoplasm as filaments. In contrast, the disordered ends of eight Nups in the nuclear ring form filaments which bind together to form a ring at the bottom of the nuclear basket. The inner ring (called the FG Nups layer in the figure above), between the cytoplasmic and nucleoplasmic rings, also consists of Nups with ordered domains and disordered parts. The disordered parts of the inner ring Nups have repetitive sequences enriched in phenylalanine and glycine (hence the name FG Nups) and stick out into the central pore. These disordered region act as a filter allowing certain molecules to pass and excluding those greater with molecular weights greater than 40K. Large molecules need transport proteins called karyopherin transport factors to move through the pore. The core structure of the NPC, obtained through cryoelectron microscopy, is shown in Figure $\PageIndex{17$ below. The double bilayers are very evident. The nuclear and cytoplasmic filaments, as well as the disordered FG repetitive sequences that stick into the pore from the inner ring, are not seen since they are very flexible and don't' adopt single conformations using standard structure determination methods. Figure $18$ shows an interactive iCn3D model of the nuclear pore complex (NPC) core (5a9q), whose structure was obtained using cryoelectron microscopy. (load slowly) A cartoon figure showing the variety of Nups in the NPC is shown in Figure $19$. In this figure, the cytoplasmic and nucleoplasmic rings are both shown in green, each mostly formed by 16 copies of the Y-complex, arranged in two eight-membered rings. The inner ring, predominantly formed by 32 copies of the Nup93 complex is shown in red. Transmembrane nucleoporins are depicted in violet, and the cytoplasmic filaments and nuclear basket structure are in orange. Attached to the inner ring are Nup62 complexes (depicted in blue), which form a cohesive meshwork within the central channel through their FG-repeat domains. Not indicated is the position of Nup98, a FG-repeat-containing nucleoporin important for the transport and exclusion function of NPCs; its position in the NPC is less defined, but it might be part of the inner ring. Similarly, Aladin (also known as AAAS), Gle1, Rae1, and Npl1 (also known as hCG1 and NUPL2) have been omitted. Large proteins and RNA that pass through the pore must first be bound to a cargo receptor, which can move the "cargo" across the pore with concomitant GTP hydrolysis. This is a process that is closer to active transport so we will discuss that in Chapter 11.3. The entire nuclear pore complex was solved in 2022 using cryoEM. Here are two videos showing the dilated complex (7R5J). Click on the images to download mp4 animations of the complex.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/01%3A_Unit_I-_Structure_and_Catalysis/11%3A_Biological_Membranes_and_Transport/11.4%3A_Diffusion_Across_a_Membrane_-_Pores.txt

Search Fundamentals of Biochemistry Introduction We have already discussed How Enzymes Work and Enzymatic Reaction Mechanisms in great detail in Chapter 6. Here we will focus on a lighter, less granular review of some key reaction mechanisms and the changes in Gibb's free energy associated with them, both in an uncatalyzed and enzyme-catalyzed reaction. Consider it a simple review of very basic organic reactions and thermodynamics in preparation for the comprehensive focus on reaction mechanisms in Unit 2: Bioenergetics and Metabolism. Breaking C-X bonds Many of the organic reactions involved in metabolism involve making and breaking bonds to carbon. There are three ways to break a bond to a C-X bond, producing either a carbocation, carbanion, or free radical intermediate, all of which are unstable and reactive, as illustrated in Figure \(1\). Both the carbocation and free radical are electron deficient, and the carbanion, although not electron deficient, has a negative on C, an atom that has a relatively low electronegativity. Figure \(1\): Ways to break a C-X bond These unstable intermediates are higher in energy than the reactants, and hence the transition state, which is even higher in energy than the intermediates, must have a structure that resembles the intermediates more than the reactants, as shown in Figure \(2\). For the charged carbanion and carbocation intermediates, there is a developing charge in the transition state. Figure \(2\): Gibb's free energy of reactants, transition state, and intermediate in breaking a bond. The thermodynamics of the reactions is determined by the change in free energy between the intermediates and the reactants, while the kinetics of the reaction is determined by the difference in free energy between the transition states and the reactants, as shown in Figure \(3\). A catalyst lowers the energy of the transition state without affecting the energies of the reactants or intermediates (assuming that these are free and not bound to the catalyst. Figure \(3\): Activation energy and ΔG for a simple uncatalyzed and catalyzed reaction The free energy diagram shown in Figure 3 is very simplistic. We need a diagram that better fits an enzyme-catalyzed reaction, using the simple reaction equation below. E + S ↔ ES → EP ↔ E + P A free energy diagram taking into account the binding of E and S followed by the conversion of bound S to bound and then the free product is shown in Figure \(4\). Figure \(4\): Simple free energy curve for the conversion for the enzyme-catalyzed conversion of substrate to product. https://commons.wikimedia.org/wiki/F...y_levels_2.svg Even this diagram is overly simplified since it suggest that the bound substrate in the ES complex is converted to the bound product in one step with no intermediates. The free energy diagram should include intermediates along the reaction pathway. An example of this is shown in Figure \(5\) for the reversible conversion of a 3-carbon sugar, dihydroxyacetone phosphate (DHAP) to another 3-carbon sugar, glyceraldehyde-3-phosphate, in a reaction catalyzed by the enzyme triose phosphate isomerase (we will study enzyme in the next chapter). Figure \(5\): Complete free energy profile for all the elementary steps of the triosephosphate isomerase catalyzed reaction. Aqvist J, Fothergill M. Computer simulation of the triosephosphate isomerase catalyzed reaction. J Biol Chem. 1996 Apr 26;271(17):10010-6. doi: 10.1074/jbc.271.17.10010. PMID: 8626554. Creative Commons Attribution (CC BY 4.0) Note that the enzyme lowers the activation energy of each of the steps in the overall reaction. Enzymes can also catalyze reactions by altering the reaction pathway although in this case, all the intermediates in the conversion are the same in both the uncatalyzed and catalyzed pathways. A comparison of the thermodynamic reactivity of molecules of similar structures can be made by determining the relative stability of the reactants and products from structural considerations. Consider two reactants, R1 and R2, which produce products P1 and P2, respectively. Any structural features that preferentially stabilize R2 compared to R1, or P2 compared to P1, but don't stabilize R2 and P2 to the same extent, will lead to a greater driving force for R2 → P2 compared to R1. This is shown graphically in Figure \(6\): Figure \(6\): Free energy reaction digrams for two similar molecules Mechanisms that lead to the stabilization of a reactant, intermediate, or product include resonance and inductive effects (electron release or withdrawal). An example of how the comparative acidity of two similar molecules can be determined through a comparison of their structures is shown below for acetic acid and ethanol is shown in Figure \(7\). Figure \(7\): Comparative acidity of two similar molecules The stronger acid, acetic acid, has the more stable (and hence less basic) charged product (the conjugate base) An example of how an intermediate can be stabilized through resonance is shown in Figure \(8\) for the keto-enol tautomerization reaction, which is favored in the direction of the keto form, a weaker acid than the enol. Figure \(8\): Comparative stability of keto and enol intermediates An example of how the inductive effect (electron release and withdrawal) stabilizes/destabilizes carbocations and cations is shown in Figure \(8\). Figure \(8\): Factors contributing to stabilization of carbocations and carbanions Also, remember that electron-withdrawing by the F's in the negatively charged conjugate base of trifluoracetic acid helps explains its lower pKa compared to acetic acid. Lastly consider how the stabilization of a tertiary carbocation below helps explain the preferential formation of the tertiary alcohol over the secondary alcohol, as shown in Figure \(9\). Figure \(9\): Preferential formation of tertiary alcohol Oxidation of Organic Molecules Organic molecules are usually oxidized in two-electron steps. Two methods can be used to determine if a C atom in an organic molecule has been oxidized. • If the number of bonds from C to oxygen increase, or the number of bonds to H decrease, the C is oxidized, More generally, if the number of bonds from C to a more electronegative atom increases, or the number of binds from C to a less electronegative atom decrease, the carbon is oxidized • A more powerful method involves determining the oxidation number of the carbon atoms in the reactant and product. If the oxidation number becomes more positive, the C is oxidized. The general rules for determining the oxidation numbers of the atoms in a molecule are: 1. O is generally 2- 2. H is usually 1+ 3. in molecules consisting of one type of atom, (like O2) - i.e. a polyatomic element, the atoms have an oxidation number of 0. 4. the sum of the oxidation numbers of the atoms in a molecule equals the net charge on the molecule or ion. In general, the oxidation number can be calculated as follows: 1. assign all nonbonded electrons of an atom to that atom 2. assign all bonded electrons to the more electronegative atom of the two atoms bonded 3. assign one electron of a bond to each atom if the two atoms are identical. 4. sum up the assigned electrons from 1-3. Subtract this number from the total number of electrons usually present in the outer shell of the atom (the group number). The result is the oxidation number. An illustration of the sequential two-step oxidation of ethane to acetic acid and assigned oxidation numbers are shown in Figure \(10\). Figure \(10\): Change in oxidation number on the stepwise conversion of ethane to acetic acid Reactions of Carbonyls: Aldehydes and Ketones When water reacts with an aldehyde in a nucleophilic addition reaction, a 1,1 diol, or a geminal diol results. This reaction can be catalyzed by a base, which acts as the nucleophile (it's a stronger nucleophile than water) and adds to the carbonyl C. OH- is regenerated when the alkoxide produced abstracts a proton from water, regenerating OH-. When an alcohol adds to an aldehyde or ketone, a hemiacetal or hemiketal, respectively, is formed. In the presence of an acid catalyst, the acid protonates the carbonyl oxygen, making the carbonyl more electrophilic. After the alcohol adds and forms the hemiacetal or hemiketal, the acid can protonate the OH group, leading to its expulsion as water in an acid-catalyzed elimination. The carbocation or resonant-form oxonium ion can react with another ROH to form an acetal or ketal. These steps are summarized in Figure \(11\). Figure \(11\): Nucleophilic addition to an aldehyde If the nucleophile is an amine, an addition can occur, followed by an elimination to form an imine or Schiff base, as shown in Figure \(12\). Figure \(12\): Schiff base formation An acetal or ketal are geminal ethers (as water addition to aldehydes or ketones produced geminal diols). As with other ethers, these geminal ethers are stable to base and are hence often used as protecting groups to keep aldehydes and ketones from undesired reactions in basic solution. Acetal formation is favored by excess anhydrous alcohol in acetic conditions, while acetal breakdown is accelerated by high concentrations of water and the presence of an acid catalyst. Why are ethers and hence acetals/ketals resistant to bases? They are resistant to nucleophilic attack, such as by base, since the expelled group (alkoxide) is unstable. (Epoxides, in contrast, will react with OH- nucleophiles since the epoxide ring is strained and of high energy.). Ethers can react with acids, however, which protonate the ether O to form an oxonium ion. Nucleophilic attack (such as by Br-) on an adjacent C can occur (SN2), with electrons flowing to the protonated oxonium ion (a great electron sink) as it departs. Reactions of Carboxylic Acid Derivatives Carboxylic acids undergo nucleophilic substitution reactions, assisted by the fact that compared to aldehydes and ketones, they have good leaving groups. With the substitution reaction, the stability of the double bond in the carbonyl is retained. Two things control the reactivity of these derivatives: the stability of the reactants and the stability of the products. The relative reactivity of carboxylic acid derivatives is shown in Figure \(13\). Figure \(13\): Relative reactivity of carboxylic acid derivatives A reactant is less reactive if stabilized by resonance. Hence the relative reactivity is as follows: amide < ester < anhydride < acid chloride The nonbonded electron pair on N of the amide, a less electronegative atom than O, can delocalize and form a resonant structure with a double bond with the carbonyl C more readily than the O in the ester. An electron pair on the bridging O in the anhydride, since its ability to form a double bond in a resonance structure, is split between the two carbonyls C is less effective in stabilizing either side than in the. (This is called competing resonances.) The reactant less stabilized by resonance is the acid chloride since a nonbonded pair of electrons on the larger chlorine molecule can't delocalize as readily given the C-Cl bond distance. Notice this order of decreased stability based on resonance stabilization is also the order of increased electrophilicity of the carbonyl C (which is most electrophilic in the absence of electron delocalization from the adjacent N, O, or Cl. The stability of the products also is important. If the deprotonated leaving group is considered as one of the products (which differentiates the different reactions), then the order of decreased stability of products is Cl- > RCOO- > RO- > RHN-.  This is shown in Figure \(14\). (Note: the pKa of ROH = 16 and R2NH = 40) Figure \(14\): Relative stability of carboxylic acid derivative leaving groups What determines the stability of products compared to reactants is the strength of bonds made and broken during the reaction. In nucleophilic addition to aldehydes and ketones, the strength of the bond to the nucleophile must be greater than the strength of the pi bond broken in the carbonyl. A C-Cl bond strength is 81 kcal/mol (340 kJ/mol) compared to a pi C-O bond strength of 93 kcal/mol (389 kJ/mol). Hence a Cl- is not likely to add to a carbonyl C. Consider the hydration of formaldehyde (carbonyl with 2 H's), acetaldehyde (with 1 H and 1 methyl group, and acetone (with 2 methyl groups). The ΔGo for hydration of these is -19, -1, and +15 kcal/mol (63 kJ/mol), respectively, showing that increased electron release toward the carbonyl C, which makes it less electrophilic and more stable, decreases the reactivity of the carbonyl. In nucleophilic substitution, the leaving group (anion) must be more stable than the nucleophile. Kinetics of Reactivity of Carbonyls The relative kinetic reactivity of various carbonyls toward nucleophiles follows the order of electrophilicity of the C. (i.e the extent of the positive charge on the carbonyl C.) The slow step in a nucleophilic attack is breaking the pi-carbonyl bond. If the reactant is stabilized by resonance in ways that reduce the electrophilicity of the carbonyl C, the reaction is slowed. Nucleophilicity is a measure of the "affinity" of an atom or ion on an electrophilic C for the nucleophilic lone pair. This is similar to basicity which is a measure of the "affinity" of an atom or ion for a proton. Halides are not good nucleophiles for reactions with acid derivatives since the halide (like Cl-) is a better leaving group than the actual leaving group. Making C-C Bonds Metabolism can be divided into catabolic (breaking down) and anabolic (synthetic) reactions. To obtain energy, sugars, and fatty acids are converted to carbon dioxide. Hence C-C bonds must be broken. In contrast, C-C bonds must be synthesized in photosynthesis. In all reactions, electrons from bond broken flow to atoms where bonds will be made. Flow is from a source (a pair of electrons possibly with a negative charge) to a sink (a slightly or fully positive atom). Figure \(15\) shows a couple of ways to make a C-C bond using either the reaction of two carbon-centered radicals (free radical mechanisms are uncommon biologically) or a carbocation with a carbanion. Figure \(15\): Making C-C bonds A carbocation is unstable unless incorporated into a molecule in which it is stable, so instead of using them, the carbonyl C is used as the electrophilic carbon. (Instability of carbocations is reflected in their propensity to rearrangement.) Taking into account the resonance form of the C=O carbonyl bond with a positive on C and a negative on O, the net charge on the carbonyl is about +0.5. A carbanion, often stabilized as an enolate resonant form, is used as the negatively charged carbon. These features are illustrated in Figure \(16\). Figure \(16\): Carbonyl carbons as electrophiles and carbanions as nucleophiles One method of making a C-C bond is an aldol condensation, in which a carbanion formed by the deprotonation of a C-H alpha to a carbonyl (which is stabilized by the enolate resonance form) acts as a nucleophile which adds to a carbonyl C in an aldehyde or ketone. The reaction is illustrated in Figure \(17\). Figure \(17\): Aldol condensation In another C-C bond synthesis reaction, a Claisen Condensation, a carbanion formed by the deprotonation of a C-H alpha to a carbonyl (which is stabilized by the enolate resonance form) acts as a nucleophile that substitutes at a carbonyl C in a carboxylic ester or thioester. This is illustrated in Figure \(18\). Figure \(18\): Claisen condensation Breaking C-C Bonds In addition to a retroaldol condensation, a common method to break a C-C is through a decarboxylation reaction at a beta-keto acid. Notice in Figure \(19\) that the analogous reaction at an alpha-keto acid is unlikely since the electrons from the C-C bond that is cleaved have no "sink" to which to flow. Figure \(19\) C-C bond cleavage by decarboxylation of beta-keto acids. Alpha-keto acids can be decarboxylate using thiamine cofactors as discussed in Chapter 6.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/12%3A_Bioenergetics_and_Biochemical_Reaction_Types/12.01%3A_Biochemical_Reactions_and_Energy_Changes.txt

Search Fundamentals of Biochemistry ATP and Phosphoryl Transfer reactions Biological oxidation reactions serve two functions, as described in the previous chapter. Oxidation of organic molecules can produce new molecules with different properties. For example, increases in solubility are observed on the hydroxylation of aromatic substrates by cytochrome P450 (which we will explore in another chapter section). Likewise, amino acids can be oxidized to produce neurotransmitters. Many biological oxidation reactions occur, however, to produce energy to drive thermodynamically unfavored biological processes such as protein and nucleic acid synthesis, or motility. Chemical potential energy is not just released in biological oxidation reactions. Rather, it is transduced into a more useful form of chemical energy in the molecule ATP (adenosine triphosphate). We will discuss the properties that make ATP so useful biologically, and how exergonic biological oxidation reactions are coupled to the synthesis of ATP. ATP is used universally as a "carrier" of free energy, which effectively means that it has a high free energy compared to its hydrolysis reaction products.  Using alternative language, ATP has a high-energy transfer potential.  The structure of ATP and a simplified reaction mechanism for the cleavage of the terminal (γ) phosphoanhydride bond by water (hydrolysis) or an alcohol (alcoholysis) is shown in Figure $1$ below. The reaction is also a nucleophilic substitution. Changing perspectives, the water or alcohol is phosphorylated by ATP. Hence the reaction can also be called a phosphoryl transfer. In Chapter 6.5, we can see that two different Enzyme Commission numbers might apply to reactions involving ATP. These are • EC2 - transferases: transfer/exchange of group from one molecule to another. More specifically they are in category EC2.7 - transferring phosphorus-containing groups • EC3- hydrolases: hydrolysis reactions. More specifically they are in category EC3.6 - acting on acid anhydrides Most of the examples in this book would best be classified as phosphoryl transfer reactions. The terminal phosphate is shown in an alternative resonance form with the P atom sp3 hybridized with tetrahedral geometry and a +1 formal charge. The attack on the electrophilic P atom by the incoming nucleophile leads to the formation of a trigonal planar transition state with the dashed lines representing bond formation and breaking in an SN2-like reaction. The final products are ADP and either inorganic phosphate (Pi) or a phosphoester. The hydrolysis reaction can be represented by a chemical equation or by a more typically written "biochemical" equation. Here is the chemical equation as recommended by the IUBMB/IUPAC: $\ce{ATP^{4-}(aq) + 2H2O (l)<=> ATP^{3-}(aq) + HPO4^{2-} (aq) + H3O^{+} (aq)} \nonumber$ This equation is written to have both charge and mass balance. For example, the sum of charges on the left-hand side (-4) is the same as on the right-hand side (-4). Of course, this is even a simplified equation since the reaction depends on the pH and the presence of divalent cations such as Mg2+. Other species that could be included just for ATP4- include HATP3, H2ATP2, MgHATP, and Mg2ATP, for example. The actual equilibrium constant would depend on the pH, the concentration of Mg2+, and the total ionic strength of the solution. If these were all fixed, the reaction could be written as a simplified biochemical equation as shown below: $\ce{ATP + H2O <=> ADP + Pi} \nonumber$ We will most often use simplified biochemical equations when discussing metabolism. Just as there are standard state conditions for chemical reactions (1 bar pressure for a gas, 1 M for a solute in solution), there are biochemical standard states for biochemical reactions. They are pressure = 1 bar, pH = 7 (i.e. H3O+=10-7 M), Mg2+ = 1 mM, and ionic strength of either 0 or 0.25 M. ATP contains two phosphoanhydride bonds (connecting the 3 phosphates) and one phosphoester bond (connecting a phosphate to the ribose ring). The pKas for the reactions HATP3- → ATP4- + H+ and HADP2- → ADP3- + H+ are about 7.0, so the overall charges of ATP and ADP at physiological pH are -3.5 and -2.5, respectively. Each of the phosphorous atoms is highly electrophilic and can react with nucleophiles like the OH of water or an alcohol. As we discussed earlier, anhydrides are thermodynamically more reactive than esters which are more reactive than amides. The large negative ΔGo (-7.5 kcal/mol, -31kJ/mol) for the hydrolysis of one of the phosphoanhydride bonds can be attributed to relative destabilization of the reactants (ATP and water) and relative stabilization of the products (ADP = Pi). Specifically • The reactants can not be stabilized to the same extent as products by resonance due to competing resonance of the bridging anhydride O's. • The charge density on the reactants is greater than that of the products • Theoretical studies show that the products are more hydrated than the reactants. The ΔGo for hydrolysis of ATP is dependent on the divalent ion concentration and pH, which affect the stabilization and the magnitude of the charge states of the reactants and products. Carboxylic acid anhydrides are even more unstable to hydrolysis than ATP (-20 kcal/mol, -84 kJ/mol), followed by mixed anhydrides (-12 kcal/mol, -50 kJ/mol), and phosphoric acid anhydrides (-7.5 kcal/mol. -31 kK/mol). The terminal anhydride bond is often called a "high energy bond".  This is absolutely wrong and has created deep misconceptions about molecules like anhydrides.   What is true is that the anhydride reactants are high energy but only compared to the energy of their cleavage products, such that the reaction proceeds with a large negative ΔGo. There is no such thing as a high-energy bond. All covalent bonds lower the energy of a system of two separated atoms. Figure $2$ shows molecules that are high energy compared to their hydrolysis products.  Think of a graph of free energy.  Carboxylic acid anhydrides and water (the reactants) have high energy compared to their collective products, two acetic acids, for example. Figure $2$: Molecules that are high energy compared to their hydrolysis products. Some older books state that the terminal anhydride bond is "high energy".  There is no such thing as a "high energy" bond.  When a bond forms between two molecules, the energy is lowered.  Energy input is required to break any bond. Each of the molecules above except the thioester has a similar motif outlined with the red dotted rectangle. The thioester also is considered high energy compared to its hydrolysis product since the reactant is effectively destabilized compared to a carboxylic acid ester. This arises because the sulfur atom is larger in the thioester than the oxygen atom in the carboxylic acid ester. Hence the bond length of C-S (1.82 Å )is larger than for C-O (1.43 Å), so the C-S bond is weaker. In addition, the lone pairs on the S, which is more polarizable than O, are less likely to be shared with the C as part of the resonance stabilization of the ester. Both effects raise the energy of the thioester compared to the carboxylic acid ester. The hydrolysis products of both esters are of similar energy. Hence the ΔGo for the hydrolysis of the thioester is more negative and about the same for the hydrolysis of ATP. How can ATP be used to drive thermodynamically unfavored reactions? First consider how the hydrolysis of a carboxylic acid anhydride, which has a ΔGo = -12.5 kcal/mol (-52 kJ/mol) can drive the synthesis of a carboxylic acid amide, with a ΔGo = + 2-3 kcal/mol (+ 4-12 kJ/mol). The reaction is: anhydride + amine --> amide + carboxylic acid This can be broken into two reactions, the hydrolysis of the anhydride, and the synthesis of the amide as shown in Figure $3$. Figure $3$: Individual and net reactions for the conversion for the formation of an amide from an anhydride and an amine Now consider the reaction of glucose + Pi to form glucose-6-P. In this reaction, a phosphoester is formed, so the reaction would proceed with a positive ΔGo = 3.3. Now if ATP was used to transfer the terminal (gamma) phosphate to glucose to form Glc-6-P, the reaction proceeds with a ΔGo = -4.2 kcal/mol (-17.6 kJ/mol). This can be calculated since ΔG and ΔGo are state functions and path independent. Adding the reactions and the ΔGos give: • glucose + Pi → glucose-6-P, ΔG0 = 3.3 • ATP + H2O → ADP + Pi , ΔG0 = -7.5 • NET: glucose + ATP -→ Glucose-6-P + ADP, ΔGo = -4.2 In most biological reactions using ATP, the terminal phosphate of ATP is transferred to a substrate using an enzyme called a kinase. Hence, hexokinase transfers the gamma phosphate from ATP to a hexose sugar. Protein kinase is an enzyme that transfers the gamma phosphate to a protein substrate. ATP is also used to drive peptide bond (amide) synthesis during protein synthesis. From an energetic point of view, anhydride cleavage can provide the energy for amide bond formation. Peptide bond synthesis in cells is accompanied by cleavage of both phosphoanhydride bonds in ATP in a complicated set of reactions that are catalyzed by ribosomes in the cells. (This topic is considered in depth in Unit III). Figure $4$ is a grossly simplified mechanism of how peptide bond formation can be coupled to ATP cleavage. Figure $4$: Individual and net reactions for the formation of a dipeptide from separate amino acids coupled with the cleavage of ATP. f In the first reaction, the carboxylic acid end of amino acid 1 is activated to form a mixed carboxylic ester. The leaving group, Pi, is hydrolyzed in reaction 2 to help drive the reaction. An amide bond is formed in reaction 3, with the expulsion of an excellent leaving group, AMP. Phosphorylation reactions using ATP are nucleophilic substitution reactions that proceed through a pentavalent transition state. These reactions are also called phosphoryl transfer reactions. One last note. ATP exists in cells as just one member of a pool of adenine nucleotides which consists of not only ATP but also ADP and AMP (along with Pi). These constituents are readily interconvertible. We break down an amount of ATP each day equal to our body weight. Likewise, we make about the same amount from the turnover products. When energy is needed, carbohydrates and lipids are oxidized and ATP is produced, which can then be immediately used for motility, biosynthesis, etc. It is very important to realize that although ATP is converted to ADP in a thermodynamically spontaneous process, the process is kinetically slow without an enzyme. Hence ATP is stable in solution. However, its biological half-life is not long since it is used very quickly as described above. This recapitulates a theme we have seen before. Many reactions (like oxidation with dioxygen, denaturation of proteins in nonpolar solvent, and now ATP hydrolysis) are thermodynamically favored but kinetically slow. This kinetic slowness is a necessary but of course insufficient condition, for life. Introduction to Active Transport We have previously discussed how chemical potential energy in the form of reduced organic molecules can be transduced into the chemical potential energy of ATP. This ATP can be used to drive reductive biosynthesis and movement (from individual cells to whole organisms). ATP has two other significant uses in the cell. Active Transport: Molecules must often move across membranes against a concentration gradient - from low to high chemical potential - in a process characterized by a positive ΔG. As protons could be "pumped" across the inner mitochondrial membrane against a concentration gradient, powered by the ΔG associated with electron transport (passing electrons from NADH to dioxygen), other species can cross membranes against a concentration gradient - a process called active transport - if coupled to ATP hydrolysis or the collapse of another gradient. Remember that active transport is different from facilitated diffusion we studied earlier, which occurs down a concentration gradient across the membrane. Many such species must be transported into the cell or intracellular organelles against a concentration gradient as illustrated in Figure $5$: Figure $5$: Examples of active transport reactions (source unfortunately lost) Signal Transduction: All cells must know how to respond to their environment. They must be able to divide, grow, secrete, synthesize, degrade, differentiate, cease growth, and even die when the appropriate signal is given. This signal invariably is a molecule that binds to a receptor, typically on the cell surface. (Exceptions include light transduction in retinal cells when the signal is a photon and lipophilic hormones which pass through the membrane.) Binding is followed by shape changes in transmembrane protein receptors which effectively transmits the signal into the cytoplasm. We will discuss two main types of signal transduction pathways: • nerve conduction, in which a presynaptic neuron releases a neurotransmitter causing a postsynaptic neuron to "fire"; • signaling at the cell surface which leads to the activation of kinases within the cytoplasm; We will discuss signal transduction in the final two chapters. For active transport to occur, a membrane receptor is required which recognizes the ligand to be transported. Of major interest to us, however, is the energy source used to drive transport against a concentration gradient. The biological world has adapted to use almost any source of energy available. Energy released by oxidation: We have already encountered the active transport of protons driven by oxidative processes. In electron transport in respiring mitochondria, NADH is oxidized as it passes electrons to a series of mobile electron carriers (ubiquinone, cytochrome C, and eventually dioxygen) using Complex 1, 3, and 4 in the inner membrane of the mitochondria. Somehow the energy lost in this thermodynamically favored process was coupled to conformational changes in the complex which caused protons to be ejected from the matrix into the inner membrane space. One can imagine a series of conformation-sensitive pKa changes in various side chains in the complexes which lead in concert to the vectorial discharge of protons. ATP hydrolysis: One would expect that this ubiquitous carrier of free energy would be used to drive active transport. This is one of the predominant roles of ATP in the biological world. 70% of all ATP turnover in the brain is used for the creation and maintenance of a Na and K ion gradient across nerve cell membranes using the membrane protein Na+/K+ ATPase. Light: Photosynthetic bacteria have a membrane protein called bacteriorhodopsin which contains retinal, a conjugated polyene derived from beta-carotene. It is analogous to the visual pigment protein rhodopsin in retinal cells. Absorption of light by the retinal induces a conformation change in the retinal and protein, which leads to the vectorial discharge of protons ; Collapse of an ion gradient: The favorable collapse of an ion gradient can be used to drive the transport of a different species against a concentration gradient. We have already observed that the collapse of a proton gradient across the inner mitochondria membrane (through FoF1ATP synthase) can drive the thermodynamically unfavored synthesis of ATP. The collapse of a proton gradient provides a proton-motive force that can drive the active transport of sugars. Likewise, a sodium-motive force can drive the active transport of metal ions. Since the energy to make the initial ion gradients usually comes from ATP hydrolysis, ATP indirectly powers the transport of the other species against a gradient. Often, the transport of one species is coupled to transport of another. If the species are charged, a net change in charge across the membrane may occur. Several terms are used to describe various types of transport, as we saw previously in Chapter 12, and which are illustrated in Figure $6$. • symport - two species are cotransported in the same direction by the same transport protein • antiport - two species are cotransported in opposite directions by the same transport protein • electrogenic - a net electrical imbalance is generated across the membrane by symport or antiport of charged species • electroneutral - no net electrical imbalance is generated across the membrane by symport or antiport of charged species

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/12%3A_Bioenergetics_and_Biochemical_Reaction_Types/12.02%3A_Phosphoryl_Group_Transfers_and_ATP.txt

Search Fundamentals of Biochemistry The History of Oxygen Oxygen may be considered one of the most important elements in chemistry. Not counting hydrocarbons, there is a greater diversity of molecules with oxygen than with carbon. Given its role in the molecular world, very little time is spent on the chemistry of oxygen in undergraduate chemistry classes. Why is oxygen so special? Oxygen reacts with atoms of all elements except the Noble gases to form molecules. One of the most important molecules of course, from a biological sense, is water. It : • provides a perfect solvent for biomolecules • moderates the earth's climate • is the source of almost all the dioxygen in the air From a chemical point of view, water is a(n): • nucleophile and electrophile • acid and base • oxidizing agent and reducing agent • a protic solvent that can form H-bonds The formation of the earth and the development of life: The gaseous and dusty environment from which the earth was formed contained metals and water, which as you remember from introductory chemistry, can react to form hydrogen gas. H2 reacts with nonmetals (under various conditions of temperature and pressure) to form H2S, HCl, CH4, and NH3 which contributed to the reducing nature of the early atmosphere. This kept the transition metals in their lowest oxidation states. Many metals, including the coinage metals (Cu, Ag, and Au) and the platinum group (Ru, Rh, Pd, Pt) were stable in elemental form. Then, around 2.7-2.8 billion years ago, photosynthetic organisms (blue/green algae- also called cyanobacteria) developed which could oxidize water to form dioxygen. Oxygen was generally unavailable for redox chemistry before then as photosynthesis, the process that would evolve to oxidize water to produce dioxygen, was unavailable. Remember that to oxidize water to dioxygen, itself a strong oxidizing agent, requires a stronger oxidizing agent than dioxygen and lots of energy. Fossilized remains of cyanobacteria are found in stromatolites. Using knowledge of how atmospheric oxygen can alter the chemistry of different sulfur isotopes of SO2, it has been shown that O2 did not exist in the atmosphere as a whole above 1 ppm earlier than 2.4 billion years ago, although there might have been isolated pockets with higher concentrations. After that, it rose, presumably as a result of cyanobacteria. Before this time, bacteria oxidized a similar molecule, H2S to form elemental sulfur. It could do this through the photosynthetic reduction of CO2 by H2S. Volcanic gases like H2 might have kept oxygen levels from rising between 2.7 billion years ago and 2.4 billion years ago when its build-up started. Hydrogen in the form of H2 and methane probably decreased around 2.4 billion years ago as methane with its hydrogen atoms escaped to the upper atmosphere and space. Methane levels would also be decreased by its easy reaction with dioxygen in the presence of UV light to form CO2. This would paradoxically lead to a cooling of the earth and pronounced glaciation as a more potent greenhouse gas, methane, was replaced with a less potent one, carbon dioxide. Over the next billion years, dioxygen rose to perhaps 0.2 - 2% (compared to the present levels of 20%) Why? Because the early atmosphere was reducing, the added oxygen combined with a large "sink" of reduced metals (like elemental Cu and Fe) or nonmetals (like C and ammonia), preventing a large buildup. Only after these reduced substances were "titrated" did dioxygen build up to present levels. In addition, the oxygen might have increased weathering (by oxidation) of sulfur deposits which can lead to sulfides entering the ocean, where they could precipitate ocean iron ions that are necessary for cyanobacterial chemistry. This would place constraints on cyanobacterial growth until dioxygen levels in the atmosphere increased enough so sulfides were converted to sulfates. This first increase in atmospheric oxygen is often called the Great Oxidation Event as it correlated and presumably caused one of the greatest mass extinctions (of anaerobic organisms) of all time. Around 2.3 billion years ago, as trace dioxygen had accumulated in the atmosphere, redox chemistry changed, although isotope evidence suggests that little dioxygen was found in water. Around 1.8 - 1.5 billion years ago, the earth's atmosphere became somewhat oxygenated, which was also coincident with the development of eukaryotic organisms. Until then, life was restricted to the oceans since there was no ozone to absorb dangerous UV radiation. The buildup of dioxygen in the air must have led to another extinction of anaerobic organisms since as we shall see, products of oxygen metabolism are very toxic. Some evolved to use dioxygen. Ozone developed, and life could then migrate from the sea to the land. It wasn't until around 600 million years ago that animals arose, however. Was this event associated with the development of a fully oxygenated (20%) atmosphere? Recent evidence, which shows that substantial oxygen wasn't available in the deep sea until about 600 million years, seems to suggest that. Based on an analysis of iron compounds in waters in Newfoundland, it appears that oxygen was very low in the sea 580 million years ago, during the Gaskier's glaciation period. Immediately after that it rose to levels consistent with atmospheric dioxygen levels of 15%, levels necessary for large animals. Similar trends in carbon and sulfur isotopes in marine rocks in Oman also suggest large increases in oxygen at the end of the Gaskiers glaciation period. What caused this second great oxygenation event? One possibility is that organic matter was sequestered from a reaction with atmospheric dioxygen, as clays bound organic molecules in the ocean and lichens and zooplankton facilitated weather and production of insoluble organic material in the oceans. Dioxygen is critically important for higher organisms, so an understanding of its chemistry becomes important. This chapter will show that dioxygen is a ground state diradical that has low solubility in an aqueous solution, reacts in a kinetically sluggish fashion in the oxidation reaction, and forms toxic byproducts as it gets reduced. Life forms hence evolved ways to deal with these problems, including ways to increase its solubility (with dioxygen binding and transport proteins), and enzymes (that could activate it kinetically and also detoxify oxygen by-products). Dioxygen is toxic to many cells. Obligate aerobes die in an oxygen environment as many of their cellular components get oxidized by this excellent oxidizing agent. Several strains of bacteria swim away from high levels of dioxygen. A graph showing the log of survival vs log pO2 is linear with a negative slope for a variety of organisms, including mice, fish, rats, rabbits, and insects. Pure oxygen can induce chest soreness, coughs, and sore throats in people. Premature infants put in pure dioxygen environments often developed blindness due to retrolental fibroplasia (a build-up of fibrous tissue behind the lens). The trade-off for this toxicity is clear. Energy is derived from organic molecules through oxidation. Before dioxygen became available to power aerobic catabolism of reduced molecules like fatty acids and less reduced sugars, such molecules were only partially oxidized. The glycolytic pathway, found in most organisms, oxidizes glucose (6 Cs) to two molecules of pyruvate (3 Cs). It was only with the availability of dioxygen did pathways evolve (Kreb Cycle, mitochondrial electron transport/oxidative phosphorylation) that allowed pyruvate to be fully oxidized to carbon dioxide, with the release of much more energy. The Properties of Dioxygen It is important to understand the properties of dioxygen since oxidation reactions using its power not only our bodies but our entire civilization. We will concentrate on biological reactions, but even these show the same characteristics as non-biological ones. • oxidation of organic molecules by oxygen is thermodynamically favored but kinetically slow. • pure oxygen environments are toxic to cells and organisms. First, we will try to understand these properties of oxygen, and then we will see how organisms overcome these problems to use dioxygen. We can understand both of these properties by looking at the molecular orbitals of oxygen and its reduction products as shown in the diagrams below. Ground state oxygen is a diradical, which explains the paramagnetic behavior of oxygen. The two unpaired oxygen each have a spin state of 1/2 for a total resultant spin S of 1, making ground state oxygen a triplet (2S+1) = 3. Organic molecules typically undergo 2 electron oxidation steps. Consider the stepwise oxidation of methane below. The oxidation number of C in methane is -4, -2 in methanol, 0 in formaldehyde, +2 in formic acid, and finally +4 in carbon dioxide, indicating two electron losses in each step. These states are shown in Figure $1$. The two electrons lost by the organic substrate are added to oxygen, but since the two lost electrons are spin paired, a spin flip must occur to allow the electrons to enter the unfilled oxygen orbitals. Alternatively, energy can be put into ground state dioxygen to produce excited state singlet oxygen (S=0, 2S+1 = 1). The source of the large activation energy required (about 25 kcal or 105 kJ/mol) to flip the electron spin accounts for the kinetic sluggishness of reactions of dioxygen with organic reactants. A traditional Lewis structure for ground state dioxygen can not be easily written since the electrons are added in pairs, and dioxygen is a diradical. There are 6 electrons in the sigma molecular orbitals from second shell electrons (two each in σ2s, σ2s*, and σ2p,) and 6 electrons in the pi molecular orbitals from second shell electrons (two each in two different π2p orbitals, and one electron each in two different π2p*), so the net number of electrons in bonding orbitals is 4, giving a bond order (or number of 2). In contrast, it is easy to write the Lewis structure of singlet, excited state oxygen, since all electrons can be viewed as paired, with two net bonds (1 sigma, 1 pi) connecting the atoms of oxygen. Figure $2$ shows the molecular orbital diagram for ground and excited state dioxygen. This Lewis structure will be used to represent singlet, excited oxygen, which should react more quickly with organic molecules. The excited state singlet on the right is unstable and decays to the middle singlet state. The middle state is approximately 94.3 kJ/mol higher in energy than the ground state triplet (on the left). In quantum mechanical parlance, the transition from the ground state triplet to the singlet state is forbidden for several reasons, making it unlikely that absorption of a photon will induce the transition. The Reduction of Dioxygen When oxygen oxidizes organic molecules, it is reduced. By adding electrons one at a time to the molecular orbitals of ground-state dioxygen we produce the step-wise reduction products of oxygen. On the addition of one electron, superoxide is formed. A second electron produces peroxide. Two more produce 2 separated oxides since no bonds connect the atoms (the number of electrons in antibonding and bonding orbitals is identical). Each of these species can react with protons to produce species such as HO2, H2O2 (hydrogen peroxide), and H2O. It is the first two reactive reduction products of dioxygen that make it potentially toxic. Figure $3$ shows the MO diagrams for the reduction products of dioxygen. How are the potential problems in oxygen chemistry dealt with biologically? Kinetic sluggishness: Enzymes that utilize dioxygen must activate it in some way, which decreases the activation energy. Enzymes that use dioxygen typically are metalloenzymes, and often heme-containing proteins. Since metals such as Fe2+ and Cu2+ are themselves free radicals (i.e. they have unpaired electrons), they react readily with ground-state oxygen which itself is a radical. The molecular orbitals of the metal and oxygen combine to produce new orbitals which for oxygen are more singlet-like. Likewise, dioxygen reacts more readily with organic molecules which can form reasonably stable free radicals, such as flavin adenine dinucleotide (FAD), as we shall see later. Dioxygen toxicity: Since toxicity arises from the reduction products of oxygen, enzymes that use oxygen have evolved to bind oxygen and its reduction products tightly (through metal-oxygen bonds) so they are not released into the cells where they can cause damage. In addition, enzymes that detoxify free dioxygen reduction products are widely found in nature. For example: • superoxide dismutase catalyzes the dismutation (self-redox) of 2 superoxides into dioxygen and hydrogen peroxide; • catalase converts hydrogen peroxide into water and oxygen; • peroxidase catalyzes the reaction of hydrogen peroxide with an alcohol to form water and an aldehyde • peroxiredoxins react with peroxides and thioredoxin (a small electron donor) to form water and oxidized thioredoxin. Finally, free radical scavengers such as vitamins A, C, E, and selenium can react with reactive free radicals to produce more stable free radical derivatives of the vitamins and Se. More on this later. The Reactions of Dioxygen and its Reduction Products Triplet O2 - Ground State Here are some reactions for the ground state (triplet O2): a. Metals ions - Metal ions are radicals themselves, so can easily react with dioxygen (think about rust). Here is one example $\ce{Fe^{2+} + O2 <=> [ Fe^{2+}-O2 <=> Fe^{3+}-O2^{-.}] <=> Fe^{3+} + \underbrace{O2^{-.}}_{superoxide}} \nonumber$ b. Autoxidation of organic molecules to produce peroxides - These are multistep reactions that have initiation, propagation, and termination steps. RH → R. (Initiation) R. + O2 → ROO. (Propagation) ROO. + RH → R. + ROOH (Propagation) R. + R. → R-R (Termination) ROO. + ROO. → ROOR + O2 (Termination) ROO. + R. → ROOR (Termination Figure $4$ summarizes the reactions of triplet ground state dioxygen. The initiation step above occurs mostly at C atoms which can produce the most stable free radicals (allylic, benzylic position, and 3o > 2o >> 10 carbons). Single O2 - Excited State Figure $5$ shows some reactions for singlet dioxygen in which dioxygen is shown as with a double bond and two lone pairs on each oxygen. Alkenes react with oxygen to form hydroperoxides, potentially through an epoxide intermediate. Dienes reacts with oxygen in a Diels-Alder pericyclic reaction to form endoperoxides. A molecular orbital perspective (that you may remember from chemistry classes) on this cycloaddition reaction is shown in Figure $6$. Singlet oxygen can be made from triplet oxygen by photoexcitation. Alternatively, it can be made from triplet oxygen through collision with an excited molecule which relaxes to the ground state after a radiationless transfer of energy to triplet oxygen to form reactive singlet oxygen. This later process accounts for the photobleaching of colored clothes when the conjugated dye molecules absorb UV and Vis light and relax to the ground state by transferring energy to triplet oxygen to form singlet oxygen. That can more readily react with the conjugated double bonds in the dye. These processes are summarized in Figure $7$. Superoxide Common reactions of superoxide are shown below. 1. Dismutation: This reaction involves a specified reactant undergoing an oxidation reaction, followed by another molecule of the same reactant undergoing a reduction. $\ce{O2^{-.} + O2^{-.} + 2H^{+} <=> H2O2 + O2} \tag{slow}$ $\ce{HO2^{.} + O2^{-.} + H^{+} <=> H2O2 + O2} \tag{fast}$ 2. Acid/Base: $\ce{HO2^{.} <=> O2^{-.} + H^{+}} \tag{pKa = 4.8}$ 3. With metal ions: $\underbrace{\ce{Fe^{3+}}}_{\text{as in heme}} + \ce{O2^{-.}} → \ce{O2} + \ce{Fe^{2+}} \nonumber$ The enzyme superoxide dismutase catalyzes the dismutation reaction. The common eukaryotic cytosolic form contains Cu2+ and Zn2+ ions, which are coordinated by histidine side chains. The reaction proceeds in two steps or half-reactions. The first is the removal (oxidation) of an electron from superoxide (O2-.) through its reduction by Cu2+. This reaction forms Cu1+ and nontoxic O2. $\ce{Cu^{2+}-SOD + O2^{−.} → Cu^{+}-SOD + O2} \nonumber$ (reduction of Cu; oxidation of superoxide) The second is the addition (reduction) of an electron from a second superoxide to Cu1+ to reform the catalytic Cu2+ and in the process form the reactive peroxide O22- (unfortunately), which when protonated forms $\ce{H2O2}$. $\ce{Cu^{+}-SOD + O2^{−.} + 2H^{+} → Cu^{2+}-SOD + H2O2} \nonumber$ (oxidation of copper; reduction of superoxide) Figure $8$ shows an interactive iCn3D model of the electrostatic potential surface of the superoxidase dismutase dimer showing Cu and Zn ions (2SOD). Two dimers are shown with a rotation C2 axis separating them. Red indicates the negative surface potential and blue the positive. Catalytic Cu2+ and Zn2+ ions are in each subunit and are shown in orange (Cu) and gray (Zn) spheres and labeled (disregard the label not centered on the orange and gray spheres). Note that the Cu2+ and Zn2+ ions are in the center of a large blue surface (positive potential), which helps "sweep up" any negatively charged superoxide O2- nearby. Rotate the complex around the C2 axis and you will see another large positive blue patch on the backside. These two positive electrostatic surfaces facilitate the electrostatic attractions and binding of the dangerous superoxide anion from the larger 3D region around the enzyme. The Zn2+ in SOD is not redox active. What is its role? Both metals appear to increase the thermostability of the individual monomer and the dimer. In the presence of metal ions (a holo form of the enzyme), the dimer dissociates into monomers at a higher urea concentration than does the apo-dimer. Ion binding reduces the flexibilities of groups found in the dimer interface. The protein has a disulfide bond between Cys 57 and Cys 146. This bond stabilizes a metal ion binding loop that contributes to the binding interface in the dimer. The enzyme operates at diffusion-controlled rates (kcat/KM is between 108 and 109 M-1s-1), in part due to the attraction of any negatively charged superoxide by the electrostatic field around the dimer. There are two other types of superoxidedismutates, one that uses either Mn2+ or Fe2+ (bacterial, mitochondrial, chloroplasts, protists) and another that uses Ni2+(some prokaryotes). Peroxide In contrast to dioxygen which contains multiple bonds between the O atoms, peroxide has only one bond. It is quite weak and requires only 38 kcal/mol (160 kJ/mol) to break it. Remember, bonds can be broken in a heterolytic way (both electrons in a bond go to one of the atoms, or in a homolytic fashion, in which one electron goes to each atom. Figure $9$ shows typical reactions of peroxides. The r​​​​eaction with Fe2+, the Fenton Reaction, is similar to the reaction of triplet O2 with Fe2+. In this reaction, homolytic cleavage of the O-O bond occurs generating OH- and the hydroxy free radical, OH., which will react with any molecule it encounters. Thermal or photochemical homolytic cleavage of peroxide also forms free radicals which react like the hydroxy free radical. The enzyme catalase facilitates the decomposition of the reactive H2O2 to water and dioxygen. As such offers protection similar to that against superoxide offered by superoxide dismutase. This is the next reaction catalyzed by the enzyme catalase: The human enzyme is a homotetramer with each monomer having a heme at the active site. The tetramer also binds NADP+ (2/tetramer) but its function is unclear. A potential mechanism for catalase is shown in Figure $10$. Figure $11$ shows an interactive iCn3D model of human erythrocyte catalase with bound ligand (CN-) and NADPH (1DGG) Figure $11$: human catalase with bound ligand (CN-) and NADPH (1DGG). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...TD7yJsfrNcwSJ7 The cyanide ligands in this structure bind to the heme in place of peroxide. The reaction in mammalian catalases appears to involve a tyrosine radical H2O2 is very similar in structure to H2O. How does the enzyme differentiate between them? Both approach the heme through a long 25 Å water channel with a hydrophobic constriction part way into the channel leading to the active site heme. Four waters in the crystal structure are positioned in the constricted opening and in a widened open just at the heme. The amino acid side chains forming the constriction are Val 74, Val 116, Pro 129, Phe 153, Phe154 and Trp 186. These allow only small molecules to enter. Two of the four waters form hydrogen bonds to side chains (one to His 75 and Asn 148 and another to Gln 168 and Asp12. The others don't form hydrogen bonds, are more dynamic, and more likely to leave the channel. H2O2 is bigger and can form bridging hydrogen bonds to side chains that the mobile waters can't. They probably leave the opening. H2O2 is more polar and has a higher dipole moment (2.26 Debye) compared to water (1.86 Debye) which implies that it would be differentially stabilized by hydrogen bonds in this hydrophobic site compared to the less polar water. Figure $12$ shows an interactive iCn3D model of human erythrocyte catalase (monomer) showing the hydrophobic constricture leading to the active site (1DGF) Figure $12$: Human erythrocyte catalase (monomer) showing hydrophobic constriction leading to the active site (1DGF). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...e48pAa9X4zhcM7 Hydroxyl Free Radical We won't specifically discuss the reaction of the hydroxyl free radical (.OH-) since it will react with anything nearby to produce another free radical. General reactions of the radical are shown in Figure $13$. In summary, we reap the benefits of using dioxygen as an oxidizing agent as it allows the aerobic and hence complete oxidation of carbohydrates and lipids to CO2 and H2O. Yet we pay the price for using O2 as an oxidizing agent as its partial reduction products, the superoxide radical, peroxides, and hydroxy free radical, collectively known as reactive oxygen species (ROS) can react with and damage proteins, nucleic acids, and lipids, as we will see below. Figure $13$ shows a summary of their reactions. We have presented the role of superoxide dismutase and catalase in the removal of ROS in cells. In addition to these elegantly designed proteins, there is another simpler biomolecule, the tripeptide glutathione (γGlu-Cys-Gly), that can reduce ROS in cells and prevent oxidative damage. Glutathione serves as a major antioxidant in cells. It exists in reduced (GSH) and oxidized (GSSG) states, the ratio of which depends on the cellular oxidative stress. It can react with and detoxify peroxides and alkyl free radicals by the following net reactions: 2 GSH + R2O2 → GSSG + 2 ROH 2GSH + 2R. → GSSG + 2RH Protection from Fe2+ - Ferritin and Transferrin The Fenton reaction shows the potential problem with having free Fe2+ ions in a chemical state that would easily allow this reaction and the generation of ROS. Hence much of the Fe2+ in the body is sequestered in Fe2+ binding cofactors like heme and FeS clusters. It is transported in the blood by the protein transferrin and stored in cells like the erythrocyte in ferritin. The iron ions in both transferrin and ferritin are in the +3 oxidation state (Fe3+). This strongly positive cation is quite insoluble in the presence of anions like hydroxide, phosphate, and carbonate. The Ksp values for the hydroxide salt of Fe3+ is 3.8x10-38. Let's detour for a bit and look at the structures of ferritin and transferrin and how they work, starting with ferritin. Ferritin The biologically functional form of ferritin is a 24-mer. The structure encapsulates a large volume that can hold many Fe ions (up to 4500) in the central cavity. The ions are stored in the more insoluble form, Fe3+, in complexes of oxide and hydroxide. Mammalian ferritin contains both heavy (H) and light (L) chains so they are hetero 24-mers. Figure $15$ shows an interactive iCn3D model of ferritin, the intracellular Fe storage protein (1fha). Figure $15$: Ferritin, the intracellular Fe storage protein (1fha). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...MQXAbPAwNnVya8 The ferritin chains are shown in purple. This structure shows only 24 Fe ions (in orange spheres with yellow halo). Two questions arise. How does Fe2+ get into the internal volume of ferritin and how does it get converted to Fe3+? There must be a channel through which Fe2+ can diffuse. Its conversion to Fe3+ requires catalysis by an Fe cluster in the heavy chain (H) subunits. Figure $16$s shows a generic diagram outlining the processes of uptake and conversion to Fe3+ salts inside the central cavity of ferritin. Heart and brain ferritin are enriched in the heavy (H) chain. These two organs clearly require safety from toxic Fe2+ ions. Ferritins in organs like the liver and spleen, which store lots of iron, are enriched in the light (L) chain. The two chains are about 50% homologous, but the H chain has a dinuclear ferroxidase iron site which catalyzes the Fe2+ to Fe3+ conversion. Once inside the L chain surface provides a nucleation site for the deposition of Fe3+ into a ferrihydrite "precipitate" ((Fe3+)2O3•0.5H2O). The general reaction is: 2Fe2+ + O2 → [Fe3+-O-O-Fe3+] → [Fe3+-O(H)-Fe3+] Figure $17$ shows a possible generic mechanism for the oxidation of the Fe cluster from Fe2+ to Fe3+. It acts as a ferroxidase that suggests that dioxygen is involved as a ligand in the oxidation of the two Fe2+ ions in the cluster to Fe3+. Figure $18$s shows a closeup of the interactions of the di-Fe cluster and two other Fe ions bound in the human H chain with water ligands. The dinuclear Fe core is shown in the central area of the figure. Figure $19$ shows an interactive iCn3D model of human heavy-chain ferritin monomer with bound Fe (4zjk) Figure $19$: Human heavy-chain ferritin monomer with bound Fe (4zjk). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?Ux6ED11dT5dyBb2XA The crystal structure of the full ferritin structure shows multiple binding sites and a channel to the oxidase site. Now let's look at how the light chain L might nucleate the formation of the iron precipitates. Crystal structures show how mineralization probably occurs at a specific site on the light chains that present themselves on the inside surface of ferritin. Figure $20$ shows a closeup of the interactions of a di-Fe cluster and two other Fe ions bound in the human L chain with water and peroxide ligands. This site probably represents the nucleation and mineralization site. Figure $21$ shows an interactive iCn3D model of human light chain ferritin with a possible nuclear site for mineralization (5LG8). The structure was made after 60 minutes of mineralization. Figure $21$: Human human light chain ferritin with a possible nuclear site for mineralization (5LG8). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...LKo7WJtoHeLBq9 The structure suggests the presence of a μ(3)-oxo)Tris[( μ (2)-peroxo)] triiron(III) cluster assembled at subsite on the L chains containing the carboxylate ligands Glu60, Glu61, and Glu64 side chains. A Glu57, which is along the incoming path of Fe ions, is involved in Fe delivery and coordination. Figure $22$ shows the electrostatic surface around the nucleation site. Note that the Fe ions are embedded in a site of negative electrostatic potential arising, in part, from the localization of the glutamic acid side chains in the site. this Why doesn't the heavy chain of ferritin perform the same nucleation function in preparation for the crystallization of ferrihydrite? A comparison of Figures 18 and 20 shows that the H chain (which has the ferroxidase activity) has a His 65 ligand instead of a glutamic acid (position 60) as one of the coordinating ligands. This gives Glu 61 more flexibility which must inhibit the nucleation and mineralization process. Figure $23$ shows two top views (left and center image) and one side view of three contiguous subunits of human heavy-chain ferritin monomer with bound Fe (4zjk). The gray/black sphere is actually a Ca2+ ion (which is larger than Fe2+) from the crystal structure. This three-monomer cluster would be replicated 8 times to form the full ferritin shell. There is a three-fold C3 axis going through the central calcium in the figure where all the monomers meet. Fe2+ must move through these central ions into the internal cavity. Transferrin Iron ions are moved in the circulation bound to the iron-binding protein transferrin. It binds to a transferrin receptor and can be endocytosed into the cell, where the Fe ions are transferred and stored in ferritin. The transferrin receptor can also bind and internalize circulating ferritin (see pdb 6GSR). Figure $24$ shows an interactive iCn3D model of transferrin (2N and 2C-lobes) binding to the ectodomain of the transferrin receptor (1SUV). Figure $24$: Transferrin (2N and 2C-lobes) binding to the ectodomain of the transferrin receptor (1SUV). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...cBhTyxoSDQgH47 The transferrin receptor is shown in gray (2 monomers) bound to the N-lobes (cyan) and C-lobes (magenta) of transferrin. Each transfer lobe has a bound CO32- (spacefill with CPK colors) and a Fe3+ ion (orange). Oxidative Modification of Proteins Many amino acid side chains can be oxidized in cells, as shown in Table $1$. Table $1$: Amino acid side chain oxidation products. Berlett and Stadtman. JBC, 272, 20313-20316 (1997). DOI:https://doi.org/10.1074/jbc.272.33.20313. Creative Commons Attribution (CC BY 4.0) Amino acids Oxidation products Cysteine Disulfides, cysteic acid Methionine Methionine sulfoxide, methionine sulfone Tryptophan 2-, 4-, 5-, 6-, and 7-Hydroxytryptophan, nitrotryptophan, kynurenine, 3-hydroxykynurinine, formylkynurinine Phenylalanine 2,3-Dihydroxyphenylalanine, 2-, 3-, and 4-hydroxyphenylalanine Tyrosine 3,4-Dihydroxyphenylalanine, tyrosine-tyrosine cross-linkages, Tyr-O-Tyr, cross-linked nitrotyrosine Histidine 2-Oxohistidine, asparagine, aspartic acid Arginine Glutamic semialdehyde Lysine α-Aminoadipic semialdehyde Proline 2-Pyrrolidone, 4- and 5-hydroxyproline pyroglutamic acid, glutamic semialdehyde Threonine 2-Amino-3-ketobutyric acid Glutamyl Oxalic acid, pyruvic acid We will focus just on one here, the oxidation of the ε-amino group of lysine by H2O2 to form α-aminoadipic semialdehyde, where the amine is replaced with an aldehyde. This reaction is called the Fenton reaction as is actually a carbonylation reaction. A hypothetical mechanism for the oxidation of lysine side chains is shown in Figure $25$. The next result is the oxidation of the lysine ε-amino group to an aldehyde. Oxidized levels of proteins (as evidenced by increased levels of carbonylation) increase dramatically with age (especially after age 40 ). The reactions seem to be catalyzed by metals and may proceed by the generation of hydroxy free radicals. Diseases associated with premature aging (Werner's Syndrome, another link to Werner's Syndrome, Progeria) show very high levels of oxidized proteins at an early age. Fibroblasts from 10 yr. old children with progeria have levels of oxidized proteins usually not seen until the age of 70. Beta-amyloid protein deposits (found in Alzheimer's and Down's Syndrome) cause neurotoxicity and death, partly by increasing superoxide production by endothelial cells, causing vasoconstriction/dilation, and ultimately disease progression. Beta-amyloid aggregates appear to increase H2O2 levels, in a process facilitated by Fe2+ and Cu+. Free radical scavengers (antioxidants) may help to prevent this damage. Carbonylation of proteins appears to be irreversible and nonrepairable. Increased carbonylation leads to misfolding and protein aggregation in ways in which protein chaperones can not reverse. The graphs in Figure $26$ (the summation of many experiments) show the correlation (negative) of increasing carbonylation of proteins (red line), a measurement of oxidative damage, and the resulting decrease in protein function (green). Red dots in the top representations of protein show carbonylation. Variants of the same protein (proteoforms) that have more intrinsically disordered regions (m1) are more susceptible to carbonylation compared to more ordered variants (m3). Cancer starts to increase around 40 years of age, and the levels of carbonylation correlate with increased cancer rates and may, in part, cause it. Lou Gehrigs Disease (Amyotrophic Lateral Sclerosis) is a disease of progressive motor neuron degeneration, which affects 1/100,000 people, and is 10-15% familial. Of the familial cases, about 25% have a mutation in superoxide dismutase I, a copper-zinc enzyme. About 2-3% of ALS patients carry 1 of 60 different dominant mutations in this enzyme. Mutations often decrease the stability of the protein which decreases Zn2+ affinity 5-50 fold. The A4V mutation (valine at amino acid 4 substituted for Ala) has the weakest Zn affinity and causes rapid disease progression. In the absence of Zn2+, the apoprotein somehow seems to induce cell death in neurons. This superoxide dismutase also expresses a second activity. It also acts as a peroxidase which takes ROH + H2O2 to form an RHO (an aldehyde) plus water. In some cases, the enzyme retains normal activity against superoxide but altered peroxidase activity. Figure $27$s shows the extent of carbonylation of wild type (WT) and two mutant forms α-synuclein after exposure (in vitro) to increasing doses of γ-radiation. α-synuclein forms aggregates (Lewy bodies) in Parkinson's Disease. The mutants, especially the A53T one, show significantly high extents of oxidative damage. This particular mutation is associated with the early onset (around 30) of Parkinson's Disease. Is your hair going white?: Wood et al have shown that millimolar concentrations of hydrogen peroxide builds up in hairs that have grayed and whitened. This was associated with a decrease in catalase and in increases in Met oxidation (to Met-sulfoxide) in proteins, also associated with a decrease in the repair enzyme Met-sulfoxide reductase, Met 374 in the active site of tyrosinase, an enzyme required for the production of melanin in hair follicles, is also damaged, leading to lack of melanin, a pigment necessary for hair coloration and "senile hair graying". ROS and Protein Folding As discussed earlier, the cytoplasm has sufficient concentrations of "β-mercaptoethanol"-like molecules (used to reduce disulfide bonds in proteins in vitro) such as glutathione (γ-Glu-Cys-Gly) and reduced thioredoxin (with an active site Cys) to prevent disulfide bond formation in cytoplasmic proteins. Disulfide bonds in proteins are typically found in extracellular proteins, where they serve to keep multisubunit proteins together as they become diluted in the extracellular milieu. These proteins destined for secretion are cotranslationally inserted into the endoplasmic reticulum (see below) which presents an oxidizing environment to the folding protein and where sugars are covalently attached to the folding protein and disulfide bonds are formed (see Chapter 3D: Glycoproteins - Biosynthesis and Function). Protein enzymes involved in disulfide bond formation contain free Cys which form mixed disulfides with their target substrate proteins. The enzymes (thiol-disulfide oxidoreductases, protein disulfide isomerases) have a Cys-XY-Cys motif and can promote disulfide bond formation or their reduction to free sulfhydryls. They are especially redox-sensitive since their Cys side chains must cycle between and free disulfide forms. Reactive oxygen species (ROS) can significantly affect redox chemistry, and if present in excess can place the cell in a condition of "oxidative" stress. ROS can indiscriminately oxidize lipids, nucleic acids, and proteins, but more specifically, they may also oxidize proteins involved in creating and maintaining the normal disulfide bond formation in proteins. As the concentration of ROS increases, the concentration of cytoplasmic proteins with incorrect disulfides should increase. Using a two dimension PAGE system (first dimension run under nonreducing and the second reducing conditions) of neural cell proteins derived from cells exposed to normal and differing oxidative conditions (hydrogen peroxide or decreased intracellular glutathione levels, Cumming et al showed that oxidizing stress increased the levels of disulfide bonds in redox sensitive enzymes and, unexpectedly, among other cytoplasmic proteins involved in many aspects of life, affecting the activity of many cellular processes, suggesting that disulfide bond formation may have not only a structural but regulatory role. Oxidative Modification of Lipids: Figure 4 shows the most likely position in organic molecules that can form stable free radicals (allylic, benzylic position, and 3o > 2o >> 10 carbons) are likely targets for reaction with ROS. Hence unsaturated fatty acids are extra reactive at the methylene C that separates the double bonds as shown in Figure $28$. Lipid and protein oxidation - cardiovascular disease The initial stages of cardiovascular disease appear to involve the development of fatty acid streaks under the artery walls. Macrophages are immune cells that have receptors that recognize oxidized lipoproteins in the blood, which they takeup. The cells then further differentiate into fat-containing foam cells which form the streaks. Oxidation of fatty acids in lipoproteins could produce lipid peroxides and along with the Fenton reaction lead to the oxidation of apoproteins in LDL. Cortical neurons from fetal Down's Syndrome patients show 3-4 times levels of intracellular reactive oxygen species and increased levels of lipid peroxidation compared to control neurons. This damage is prevented by treatment of the neurons in culture with free radical scavengers or catalase. Key events in atherosclerotic plaque initiation are shown in Figure $29$. How do fatty streaks appear under the endothelial cells? LDL oxidized in the lipid monolayer and through carbonylation of lysine side chains of the apoproteins in LDL binds to "scavenger" receptors in macrophages which have moved into the intima below the endothelial cell barrier. Scavenger receptors were first recognized to bind acetylated LDL (conversion of ε-amino groups of lysines, for example, to acetylated and uncharged derivatives). This mimics to some degree the carbonylation of the ε-amino groups to aldehydes, as shown in Figure $30$. Either modification would make the apoprotein more acidic with a lower isoelectric point since positive lysine side chains are replaced with neutral derivatives. Assuming an asymmetric distribution of the negatively charged side chains (Asp and Glu) on the apoprotein, any pre-modification negative electrostatic potential surfaces on the apoprotein would become more negative, enhancing binding to positive clusters displaying positive electrostatic potentials on scavenger receptors. Scavenger receptors often bind polyanions. There are many 12 different classes (A-L) of scavenger receptor classes that have been identified. One, class C, is only found in drosophila. They bind a variety of polyanionic ligands and display broad binding specificity. Many in a single class have multiple names that makes their designation even more confusing. They bind a diverse set of ligands including those from bacteria and yeast (in a way similar to pathogen-associated molecular patterns - PAMPs - in the innate immune system) as well as self and modified self ligands (such as oxidized LDL - oxLDL). Once bound the ligand and scavenger receptors are taken into the cell by endocytosis for removal and degradation of the bound ligand. They can also act in signaling pathways. The members of the scavenger receptor family are designated as illustrated in this example, SR-F1.1, where S is Scavenger, R is Receptor, F is Class, 1 is Order in class and 1 is alternatively spliced forms. Figure $31$ shows domain structures of the different classes of scavenger receptors. Figure $31$: Domain structures of the different classes of scavenger receptors. Zani et al. Cells 2015, 4, 178-201. https://doi.org/10.3390/cells4020178. Creative Commons Attribution 4.0 International A more detailed cartoon showing examples from a few different scavenger receptor classes involved in cardiovascular disease is shown in Figure $32$. Modified LDL binds to several different scavenger receptors, including SR-A1 (also called SCARA1 or CD204), SR-A2 (also called MARCO), and SR-E1 (also called Lectin-like oxidized LDL receptor 1 or LOX-1) binds oxidized and acetylated LDL. Let's look in greater detail at two scavenger receptors that recognized oxLDL SR-A2 (MARCO) This scavenger receptor is a trimer that binds oxLDL, polyanions, and pathogens. It has an extracellular domain (ectodomain) formed from three monomers that are cysteine rich, so it's abbreviated SRCR. A five-stranded antiparallel β-sheet, and an α-helix with a large loop covering it, while the dimer has a larger 8-stranded eight-stranded β-sheet. The polyanion ligands bind presumably to the surface of the receptors with a positive (blue in figures) electrostatic potential associated with an arginine cluster. Crystal structures show that the protein also has a region of negative (red in figures) electrostatic potential which most likely is involved in metal ion binding and in the self-association of monomers to form trimeric receptors. Given the size of the oxidized LDL (250 Å in diameter), it would not be unexpected that oxLDL binding would promote the formation of clusters of the normally trimer scavenger receptor. This is illustrated in Figure $33$. Monomeric, dimeric, and oligomeric forms of SR-A2 (MARCO) are shown in the bottom part of the figure. The trimeric receptor molecules can form dimers and multimers by swapping domains. Multiple interactions would promote tighter binding of large ligands such as LDL and even bacteria (0.2-2 μm diameter). The assembly could proceed to the formation of oligomers (the yellow molecule has swapped domains with three other molecules), thus resulting in the creation of a large surface capable of interacting with large ligands, such as modified LDL (250 Å in diameter) or bacteria (0.2-2 μm). The red and blue surfaces shown above the trimer represent the negative (red) and positive (blue) electrostatic surface electrostatic potential of the oligomeric from top down. SR-E1 (Lox1 or Lectin-like oxidized LDLR or Oxidized low-density lipoprotein receptor 1. LOX-1 is expressed on macrophages, dendritic cells, endothelial cells, platelets, smooth muscle cells, and adipocytes. It binds oxLDL, some bacteria (through their negatively charged cell walls), and even apoptotic cells. Figure $34$ shows an interactive iCn3D model of the extracellular C-type lectin-like domain of dimeric human Lox-1 (1YPQ) Figure $34$: Extracellular C-type lectin-like domain of dimeric human Lox-1 (1YPQ). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...qGY4HBzPjjs7P7 The dimer in this structure is connected by a disulfide bridge. The two monomers are shown in gray spheres. They come together to form a heart-like structure A series of arginines (with blue spheres for the surface Ns and labeled) are shown in the cradle of the heart shape. These most likely interact with the oxidized apoprotein B of the oxLDL. Other blue (N) and red (O) spheres near to each other can form salt bridges and may interact with zwitterion heads of LDL surface lipids such as phosphatidylcholine, sphingomyelin, or phosphatidylethanolamine. Figure $35$ shows an interactive iCn3D model of an AlphaFold predicted model of human oxidized LDL receptor - LOX (P78380) Figure $35$: . (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...rLoi7ZK4K3SsH8 The yellow spacefill indicates the transmembrane segment. Rotate the model along the long axis and you will see one face of the top domain has a red (negative electrostatic potential) face while the opposite side has blue (positive potential). Oxidative Modification of DNA Significant evidence suggests oxygen free radicals are linked to aging and diseases. Mutations caused by hydroxylation reactions (presumably from the generation of hydroxyl free radicals as shown above) can potentially lead to cancer. A particularly nasty reaction is the insertion of the hydroxy radical into bases in DNA. Figure $36$ shows the hydroxylation at position 8 of guanine to produce 8-oxy-G and at positions 5 and 6 in thymine. Mitochondrial DNA is more susceptible to oxidation than is nuclear DNA. The human mitochondrium has a small genome (16.5 Kb compared to the nuclear genome of 3 Gb) which code 13 protein subunits involved in respiration, 22 tRNAs and two ribosomal RNAs. (The mitochondria presumably are vestiges of a bacteria which invaded an early cell and established a symbiotic relationship with the cell). There is an inverse correlation of oxidized mitochondrial DNA [8-oxoG] with the maximal life span of an organism, but this correlation is not seen with nuclear DNA. Presumably,  the nuclear DNA is somewhat protected from oxidative damage since it is bound to histone proteins (which form nucleosome core particles with DNA) and by DNA repair enzymes. DNA repair enzymes that are encoded in the nucleus are found in the mitochondria and mitochondrial DNA is packaged with mitochondrial transcription factor A (TFAM). Examination of human bladder, head and neck, and lung primary tumors reveals a high frequency of mitochondrial DNA mutations. In addition,  most dioxygen use by the cell occurs in the mitochondria. Hence this organelle probably faces the highest concentration of toxic oxygen reduction products. Recently, the crystal structure of an enzyme, adenine DNA glycosylase (MutY), that repairs 8-oxyG modified DNA has been determined in complex with the oxidatively damaged DNA. If not repaired, the 8-oxyG base pairs with adenine instead of cytosine, causing a GC to AT mutation on DNA replication. Figure $37$ shows an interactive iCn3D model of adenine mispaired with 8-oxoguanine by MutY adenine DNA glycosylase (1RRQ) The protein MutY, which catalyzes the base excision and repair, is shown in gray. The DNA strands are shown in magenta and blue. 8-OxyG on the magenta strand is labeled 8OG7 and is shown in CPK-colored sticks. Its mismatched adenine base pair partner, labeled A18 on the blue strand, is shown in CPK-colored sticks. Notice its orientation is kinked away from the orientation in a canonical base pair. Key amino acid side chains (Thr49, Leu86, Tyr88, and Ser308) interacting with the 8-oxyG are shown in CPK-colored sticks and labeled. Although oxidative damage in mitochondria clearly can promote premature aging, other independent mechanisms may also. Kujoth et al. developed a mouse model that expressed a mutant form of mitochondrial DNA polymerase that was defective in the proofreading activity of the enzyme. These mice displayed premature aging but showed no increased levels of oxidized mitochondrial lipids or hydroxylated G residues in mitochondrial DNA. They did show significant activation of a cytosolic enzyme called caspase-3, which when active lead to the programmed death of cells (a process called apoptosis). This calcium-activated aspartic acid protease (with an active site Asp) is activated by binding mitochondrial cytochrome C that has "leaked" into the cytoplasm from its normal location in the intermembrane space in mitochondria. The process is usually associated with DNA damage (mutations, fragmentation) that would arise if the proofreading function of DNA polymerase was defective. This was indeed found in these mice. Oxidative damage to biomolecules might not initiate aging and disease processes, but rather might be a secondary effect of other initiating events. .Reversing or preventing oxidative damage might slow the progression of aging and disease. Aging is a complex feature of organisms and would be expected to have complex causes and biological effects. At the organismal level, aging has been studied in the roundworm C. elegans which lives for only a few weeks. Genetic analyses can be easily used to find gene alterations associated with premature aging. One hormonal system that has recently been associated with aging in eukaryotes (and in C. elegans) involves the signaling pathways for insulin and insulin growth factor I (IGF-1), which regulate carbohydrate, lipid, and reproductive pathways in C. elegans. Mutations that decrease signaling from this pathway increase C. elegans life span. These mutations lead to increased activity of the DAF16 transcription factor, which upregulates the expression of many genes. In contrast, wild-type organisms, when exposed to insulin or IGF-1, decrease the activity of DAF16. Using DNA microarrays, investigators determined which DAF16-controlled genes were upregulated in mutant worms in the mid-life point of the organism. These genes included, among others, peroxisomal and cytosolic catalase, Mn-superoxide dismutase, cytochrome P450s, metallothionein-related Cd-binding protein, and heat shock proteins. We will investigate the function of several of these gene products in the next section, but needless to say, they are all involved in cellular responses to stress, often involving dioxygen metabolites. The over-expression of mitochondrial catalase in mice increased their lifespan by 20%. It has also been showed that decreased levels of insulin-like growth factor also promote longevity in mice, indicating again that mechanisms in addition to oxidative damage by ROS are involved in aging. Beneficial Oxidation of Proteins: Oxidative Burst in Macrophages There are cases in which oxidative damage to protein and lipids is desirable. One example involves the role of macrophages in the immune system in eliminating foreign microorganisms. When macrophages recognize and engulf microbes, one mechanism deployed in killing the microorganism is through oxidative damage. The stimulated macrophages undergo an oxidative burst which leads to increased oxygen utilization. One outcome of this is the generation of ROS. The activation of the ROS-generating system can also kill the macrophage (which is OK). In addition, immune function decreases with age. This probably also occurs through damage from ROS. Telomeres at the end of chromosomes are also shortened by oxidative stress and irradiation. They are enriched in guanine bases the have may repeat (thousands) of TTAGGG sequences and hence are susceptible to oxidation. However, as you might expect, macrophages have developed ways to limited self-damage by ROS. In the presence of ROS, the macrophage or mitochondrial kinase Mst1/2 are recruited to their respective membranes from the cytosol. The enzyme acts as a ROS sensor and "attenuator" though the phosphorylation and stabilization of a protein Keap1 that binds to a transcription factor Nrf2 (also called NFE2L2), a transcription factor. When bound to unphosphorylated Keap1, the transcription factor Nrf2 is target for proteolysis. Keap1 phosphorylation prevents its binding to Nrf2. Free Nrf2 then can translocate to the nucleus where it promotes transcription of antioxidant proteins such as glutamate-cysteine ligase catalytic subunit (Gclc), which catalyzes the first step and rate-limiting step of glutathione (g-glutamyl-cysteinyl-glycine) synthesis. These processes are illustrated in Figure $38$. Phagosomal or mitochondrial ROS release attracts Mst1/2 to the membrane of phagosome or mitochondrion from the cytosol and activates Mst1/2; Mst1/2 phosphorylate Keap1 to stabilize Nrf2 and regulate the expression of antioxidant enzymes to protect the cell against oxidative damage.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/12%3A_Bioenergetics_and_Biochemical_Reaction_Types/12.03%3A_The_Chemistry_and_Biochemistry_of_Dioxygen.txt

Search Fundamentals of Biochemistry General Oxidizing Agents Before we consider common biological oxidizing agents, lets look back at ones you saw in other chemistry classes. Oxidizing agents are required to oxidize organic molecules. In organic lab, you never used dioxygen as an oxidizing agent. It is difficult to limit the extent of oxidation using dioxygen. In addition, side reactions are likely given the nature of the reactive oxygen reduction products. The mechanisms of combustion reactions of organic molecules with dioxygen to produce carbon dioxide and water are very complicated. Table $1$ belows some key steps in the combustion of methane. Initiation $\ce{CH4 → CH3^{.} + H^{.}} \nonumber$ $\ce{O2 → 2 O^{.}} \nonumber$ Propagation $\ce{CH4 + H^{.} → CH3^{.} + H2} \nonumber$ $\ce{CH4 + HO^{.} → CH3^{.} + H2O} \nonumber$ $\ce{CH3^{.} + O^{.} → CH2O + H^{.}} \nonumber$ $\ce{CH2O + HO^{.} → CHO^{.} + H2O} \nonumber$ $\ce{CH2O + H^{.} → CHO^{.} + H2} \nonumber$ $\ce{CHO^{.} → CO + H^{.}} \nonumber$ $\ce{CO + HO^{.} → CO2 + H^{.}} \nonumber$ Branching $\ce{H^{.} + O2 → HO^{.} + O^{.}} \nonumber$ Termination $\ce{H. + R^{.} + M → RH + M^{*}} \nonumber$ In chemistry, other oxidizing agents are often used, including permanganate and chromate. Their mechanism of action is illustrated in Figure $1$. Oxygen can often be inserted into a molecule in a nonoxidative process by hydration of an alkene to an alcohol (a readily reversible reaction), which could then be oxidized to either an aldehyde/ketone or carboxylic acid using an appropriate oxidizing agent. Most biological oxidation reactions (such as those found in glycolysis, Kreb Cycle, and fatty acid oxidation) do not use dioxygen as the immediate oxidizing agent. Rather they use nicotinamide adenine dinucleotide (NAD+) or flavin adenine dinucleotide (FAD) as oxidizing agents, which get reduced. Enzymes that uses these oxidizing agents are usually called dehydrogenases. Dioxygen can also be used to introduce oxygen atoms into biological molecules in oxidative reactions. Enzymes that introduce one oxygen atom of dioxygen into a molecule (and the other oxygen into water) are called monooxygenases. (Note: some monooxygenase that hydroxylate biomolecules are called hydroxylases.) Those that introduce both atoms of dioxygen into a substrate are called dioxygenases. These oxygenases are not usually used to oxidize organic molecules for energy production. Rather they introduce O atoms for other reasons, including increasing the solubility of nonpolar aromatics to facilitate secretion, and to produce new molecular species which have different biological activities. Finally, biological molecules can be oxidized by dioxygen in which no atoms of oxygen are added to the substrate. Rather, electrons lost from the oxidized substrate are passed via intermediate electron carriers to dioxygen , which get reduced to superoxide (if one electron is added), hydrogen peroxide (if two electrons are added) or water (if 4 electrons are added). These enzymes are called oxidases. (Note: The letters oxi- or oxygen- are used in all the enzymes that use dioxygen as the oxidizing agent.) In this chapter section, we will discuss biological oxidation reactions. Most introductory biochemistry texts don't approach oxidation reactions in one cohesive chapter. Probably because of that, when I was learning biochemistry, I found the presentation of these different enzymes involved in redox reactions to be very confusing. Hopefully this section will alleviate that problem. First the chemistry of NAD+ and FAD will be discussed. Then the enzymes using dioxygen in oxidative reactions (monooxygenases, dioxygenases, and oxidases) will be explored. The Chemistry of NAD+ and FAD NAD+ is a derivative of nicotinic acid or nicotinamide, as illustrated in Figure $2$. Figure $2$: Structure of niacin derivatives It and its reduction product, NADH, exists in the cells as interconvertible members of a pool whose total concentration does not vary significantly with time. Hence, if carbohydrates and lipids are being oxidized by NAD+ to produce energy in the form of ATP, levels of NAD+ would begin to fall as NADH rises. A mechanism must be be present to regenerate NAD+ from NADH if oxidation is to continue. As we will see later, this happens in the muscle under anaerobic conditions (if dioxygen is lacking as when you are running a 100 or 200 m race, or if you are being chased by a saber-toothed tiger) when pyruvate + NADH react to form lactate + NAD+. The reaction is shown in Figure $3$. Figure $3$: Conversion of pyruvate to lactate Under aerobic conditions (sufficient dioxygen available), NADH is reoxidized in the mitochondria by electron transport through a variety of mobile electron carriers, which pass electrons to dioxygen (using the enzyme complex cytochrome C oxidase) to form water. NAD+/NADH can undergo two electron redox steps, in which a hydride is transferred from an organic molecule to the NAD+, with the electrons flowing to the positively charged nitrogen of NAD+ which serves as an electron sink. NADH does not react well with dioxygen, since single electron transfers to/from NAD+/NADH produce free radical species which can not be stabilized effectively. All NAD+/NADH reactions in the body appear to involve 2 electron hydride transfers. Figure $5$ shows both 1 and 2 electrons to NAD+. Figure $4$: Figure $4$: 1 and 2 electrons to NAD+ FAD (or flavin mononucleotide-FMN) and its reduction product, FADH2, are derivatives of riboflavin, as shown in Figure $5$. Figure $5$: Structures of riboflavin, FMN and FAD FAD/FADH2 differ from NAD+/NADH since they are bound tightly (KD approx 10-7 - 10-11 M) to enzymes which use them. This is because FADH2 is susceptible to reactions with dioxygen, since FAD/FADH2 can form stable free radicals arising from single electron transfers. FAD/FADH2 can undergo 1 OR 2 electrons transfers. This is illustrated in Figure $6$. Figure $6$: 1 and 2 electrons reduction of FAD FAD/FADH2 are tightly bound to enzymes so as to control the nature of the oxidizing/reducing agents that interact with them. (i.e. so dioxygen in the cell won't react with them in the cytoplasm.) If bound FAD is used to oxidize a substrate, the enzyme would be inactive in any further catalytic steps unless the bound FADH2 is reoxidized by another oxidizing agent. Dehydrogenases These enzymes use NAD+/NADH or FAD/FADH2 and are named for the substrate that is oxidized. For instance in the reaction: $\ce{pyruvate + NADH <=> lactate + NAD^{+}} \nonumber$ which is used to regenerate NAD+ under anerobic conditions, the enzyme is named lactate dehydrogenase. As in acid/base reactions, when the preferred direction for the reaction (from a ΔGo perspective) is from stronger acid to weaker (conjugate) acid, the preferred direction for a redox reaction is in the direction from strong to weak oxidizing/reducing agents. This can easily be determined from charts of standard reduction potentials, and using the equation: ΔGo = -nFEo, • where F is the Faraday constant (96,494 Coulombs/mol e- = 96, 494 J/(V.mol) = 23.06 kcal/(V.mol) or 96 kJ/(V.mol). One Faraday is the charge per one mol of electrons). • and Eo, the standard EMF or standard cell potential (total voltage at standard state conditions), which can be determined by adding the standard reduction potentials (Eo) for the two appropriate half-reactions, after reversing the equation for the half-reaction that represents the oxidation. Hence Eo=Eoreduction−Eooxidation. When n=2 (number of electrons) which is common for oxidations of organic molecules, ΔGo (kcal/mol) = - 46.12Eo or approximately -50E0  (or -193Eo kJ/mol) Notice when Eo > 0, ΔGo < 0, the reaction as written is favored under standard conditions. Note in the table below that many of the half reactions involve protons. For biological reactions involving free protons, the standard state concentration for the protons are not 1 M as for other solutes in solution, but defined to be the hydronium ion concentration at pH 7.0. The Eo and ΔGo values for the reactions involving hydrogen ions at a standard state of pH 7.0 are usually written as Eo' and ΔGo'. Common standard reduction potentials are shown in Table $2$ below. Table $2$: Standard reduction potentials for common biomolecules oxidant reductant n (electrons) Eo' (volts), 25oC Acetate + carbon dioxide pyruvate 2 -0.70 succinate + CO2 + 2H+ α−ketoglutarate + H2O 2 -0.67 acetate acetaldehyde 2 -0.60 glycerate-3-P glyceraldehyde-3-P + H2O 2 -0.55 O2 O2- 1 -0.45 ferredoxin (ox) ferredoxin (red) 1 -0.43 Carbon dioxide formate 2 -0.42 2H+ H2 2 -0.42 α-ketoglutarate + CO2 + 2H+ isocitrate 2 -0.38 acetoacetate β-hydroxybutyrate 2 -0.35 Cystine cysteine 2 -0.34 Pyruvate + CO2 malate 2 -0.33 NAD+ + 2H+ NADH + H+ 2 -0.32 NADP+ + 2H+ NADPH + H+ 2 -0.32 FMN (enzyme bound) FMNH2 2 -0.30 Lipoic acid, ox Lipoic acid, red 2 -0.29 1,3 bisphosphoglycerate + 2H+ glyceraldehyde-3-P + Pi 2 -0.29 Glutathione, ox Glutathione, red 2 -0.23 FAD (free) + 2H+ FADH2 2 -0.22 Acetaldehyde + 2H+ ethanol 2 -0.20 Pyruvate + 2H+ lactate 2 -0.19 Oxalacetate + 2H+ malate 2 -0.17 α-ketoglutarate + NH4+ glutamate 2 -0.14 FAD + 2H+ (bound) FADH2 (bound) 2 0.003-0.09 Methylene blue, ox Methylene blue, red 2 0.01 Fumarate + 2H+ succinate 2 0.03 CoQ (Ubiquinone - UQ) + H+ UQH 1 0.031 UQ + 2H+ UQH2 2 0.06 Dehydroascorbic acid ascorbic acid 2 0.06 Ubiquinone; ox red 2 0.10 Cytochrome b2; Fe3+ Cytochrome b2; Fe2+ 1 0.12 Cytochrome c1; Fe3+ Cytochrome c1; Fe2+ 1 0.22 Cytochrome c; Fe3+ Cytochrome c; Fe2+ 1 0.25 Cytochrome a; Fe3+ Cytochrome a; Fe2+ 1 0.29 1/2 O2 + H2O H2O2 2 0.30 Cytochrome a3; Fe3+ Cytochrome a3; Fe2+ 1 0.35 Ferricyanide ferrocyanide 2 0.36 Cytochrome f; Fe3+ Cytochrome f; Fe2+ 1 0.37 Nitrate nitrite 1 0.42 Photosystem P700 . . 0.43 Fe3+ Fe2+ 1 0.77 1/2 O2 + 2H+ H2O 2 0.816 The mechanism for the oxidation of a substrate by NAD+ involves concerted hydride transfer to one face of NAD+. Hydride transfer is possible since water is excluded from the active site. It is facilitated by removal of a proton from an oxygen on an alcoholic substituent, for example, adjacent to the departing hydride. The negative charge on the oxide acts as a "source" of electrons, which can then flow through the hydride transfer to the positively charged ring nitrogen of NAD+, which acts as an electron "sink". This is crudely illustrated in the cartoon shown in Figure $7$, which shows the oxidation of ethanol by the enzyme alcohol dehydrogenase. For substrates like ethanol that lose a hydride from a methylene carbon atom that has two hydrogens, only one of them is lost (either the proR or proS) from the prochiral center. The site on NAD+ that receives the hydride, as well as the entire ring, is planar with sp2 hybridization. When bound to the enzyme, the hydride is transfer to the Re face of he ring. The same occurs in the reverse reaction when the hydride from NADH is transferred to the re face of acetaldehyde. Re and Si faces can be determined using by prioritizing the substituents attached to the sp2 carbon using the Cahn-Ingold rules. The reversible transfer of the proR hydrogens to the Re faces of the reactants in the reversible conversion of ethanol to acetaldehyde are shown in Figure $8$. Figure $8$: Reversible transfer of the proR hydrogens to the Re faces of the reactants in the alcohol dehydrogenase reaction FAD has a more positive reduction potential than NAD+ so it is used for more "demanding" oxidation reactions, such as dehydrogenation of a C-C bond to form an alkene. You will notice on standard reduction potential tables that the potential of FAD is often listed several times and depends on the enzyme. This is because the FAD is tightly bound to the enzyme so its tendency to acquire electrons depends on its environment, in much the same fashion as the pKa of an amino acid side chain (which reflects is tendency to release protons) is affected by the environment of the amino acid side chain in the protein. The standard reduction potential for flavin enzymes varies from -465 mV to + 149 mV. Compare this to the reduction potential of free FAD/FADH2, which in aqueous solution is -208 mV. The standard reduction potential of the flavin in D-amino acid oxidase, a flavoprotein, is about 0.0 V. Remember, the more positive the standard reduction potential, the more likely the reactant will be reduced and hence act as an oxidizing agent. Hence the FAD in D-amino acid oxidase is a better oxidizing agent than free FAD. The KD for binding of FAD to the enzyme is 10-7 M compared to the KD for binding of FADH2, which is 10-14 M. By gaining electrons, the flavin binds more tightly, which preferentially stabilizes the bound FADH2 compared to the bound FAD. This shifts the equilibrium of FAD ↔ FADH2 to the right, making the bound FAD a stronger oxidizing agent. A mechanism for the 2-electron hydride reduction of FAD is shown in Figure 6 above. Figure $9$ below shows an interactive iCn3D model of D-amino acid oxidase bound to FAD and a trifluoroalanine (1C0L) . Figure $9$: D-amino acid oxidase bound to FAD and a trifluoroalanine (1C0L). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...wYfRAK9TBSt2c8 FAD is shown in spacefill and colored yellow. Only the amino acids interacting with FAD are shown as a surface with underlying CPK colored sticks. The noncovalent interactions are shown as sticks. Note that you rotate the molecule, you will see the the FAD is almost completely buried which, along withe the extensive interactions with protein contributes to its low KD. Can the standard reduction potential of a redox active center in a protein be tuned by changing the environment of that center, much as the pKa of an acid side chain can by changing the polarity of the environment? The answer is yes. The active site of azurin, a cupredoxin, has a redox active copper ion coordinated by a Cys and two His residues in a trigonal planar fashion. Met 121 serves as a weak axial ligand. Marshall et al. have reported a feasible method to manipulate the redox potential (Eo) of this active site. The wild type azurin was mutated to alter the hydrophobicity and hydrogen bonding capabilities, while maintaining the overall architecture of the metal binding site. Ser 46 was selected for mutation since it occupied a position similar to Asn in another cupredoxin that was involved in an important H bond binding two ligand binding loops. An N47S-mutation, which strengthened the hydrogen bond between the two ligand-containing loops increased Eo by ~130 mV while preserving metal binding site architecture as determined by UV-Vis spectroscopy. They also compared a M121Q mutant with wild-type M121 and with a M121L mutant. A plot of Eo vs log partition coefficient for transfer of the side chain from water to octanol was essentially linear with a positive slope, showing that the standard reduction potential depended on the hydrophobicity of the weakly coordinating ligand in the metal binding region. This behavior extended to double mutants (where one set of mutants involved M121). The investigators were able to tune the Eo over a 700 mv range! Monooxygenases An examples of monooxygenases are the hydroxylases which hydroxylate amino acids like tryptophane and tyrosine to form 5-hydroxytryptophan and 3-4-dihydroxyphenylalanine or dopa, respectively. These latter two substances can be decarboxylated using PLP-dependent enzymes to form the neurotransmitters 5 hydroxytryptamine (5HT or serotonin) and dopamine. The latter can be hydroxylated again to form norepinephrine, and subsequently methylated to form epinephrine. LSD and amphetamine are analogs of serotonin and dopamine, respectively. The derivative of tryptophan and tyrosine are shown in Figure $10$. Figure $10$: Derivative of tryptophan and tyrosine derived from monooxygenases Since these monooxygenases use dioxygen, you might expect that the enzymes would use the motifs described in the previous section to facilitate its reaction with dioxygen. In fact, the enzyme contains a metal ion (Fe2+) bound to a heme in the protein. In addition, the reduction products of dioxygen that are eventually used to hydroxylate the substrate stay bound to the enzyme. Tyrosine 3-monooxygenase (Tyrosine hydroxylase) Tyrosine hydroxylase is a homotetramer which uses an Fe ion and biopterin cofactor for hydroxylation. In the central nervous system is used in the biosynthesis of dihydroxyphenylalanine (DOPA), which is the rate limiting step in the catecholamine synthesis. A possible mechanism for the rat enzyme is shown in Figure $11$. Figure $11$: A possible mechanism for rat tyrosine hydroxylase, a monooxygenase. https://www.ebi.ac.uk/thornton-srv/m-csa/entry/134/ The Fe2+ ion binds O2. Some mechanisms show a single electron transfer to the bound O2 from the pterin ring to form Fe2+-O2-(bound superoxide) and a radical cation pterin. This is unstable and forms Fe2+-μ-peroxypterin, followed by heterolytic cleaves of the peroxo O-O bond. Ultimately, hdroxpternin and an Fe4+O oxospecies form which then hydroxylates tyrosine. Figure $12$ below shows an interactive iCn3D model of rat tyrosine hydroxylase with bound cofactor analogue and iron (2TOH). Figure $12$: Rat tyrosine hydroxylase with bound dihydrobiopterin analogue and iron (2TOH). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...KeKiLDfmeNFfr7 Four monomers in the homotetramer are shown. The cofactor analog (HBI), 7,8-dihydrobiopterin, is shown in spacefill. Phenylalanine 300 has been self-hydroxylated by the enzyme to produce meta-tyrosine (MTY300), which along with the other key amino acids in the active site are shown labeled in CPK-colored sticks in the gray monomer. The pterin ring interacts through pi stacking with Phe 300, which facilitates self-hydroxylation. The Fe ion is far (5.6 Å) from the carbon on pterin which gets phosphorylated. This suggests that O2 might bridge the reacting pterin C- and the Fe ion in a Fe2+-μ-peroxypterin complex. Tryptophan hydroxylase This enzyme also uses a tetrahyropterin cofactor and Fe ion in the formation of he hydroxylating intermediates As in the case of tyrosine hydroxlase, both the amino acid substrate and the cofactor must be present for oxygen to be activated. Other, it only oxidizes Fe2+ to Fe3+. This serves as a protective mechanism so that O2 is not activated unnecessarily, which avoids the formation of soluble ROS. Figure $13$ shows a possible mechanism without catalytic residues for tryptophan hydroxylase Figure $13$: possible mechanism without catalytic residues for tryptophan hydroxylase adapted from Kenneth M. Roberts, Paul F. Fitzpatrick, https://doi.org/10.1002/iub.1144 Cytochrome P450s The cytochrome P450 (CYP)consists of a large group of monooxygenases that contain a heme that absorbs maximally at 450 nm. They catalyze the following reaction: RH + NAD(P)H + H+ + O2 → ROH + NAD(P)+ + H2O An example includes cytochrome P450cam that hydroxylates camphor, a large aromatic completely nonpolar molecule. The enzyme has been called the"biological equivalent of a blowtorch" as it can, at room temperature, stereospecific hydroxylate nonactivated hydrocarbons at physiological temperature. This reaction proceeds without stereospecificity only at high temperatures in the absence of a catalyst. Remember from the previous chapter section that dioxygen is a ground state triplet radical which can't react well with carbons atoms (which are singlets) unless converted to a singlet state, a process which requires significant energy. Cytochrome P450s get around this problem by binding to an Fe ion in the heme to form common intermediates such as oxos, oxides, and peroxides. The molecular orbitals of the bound dioxygen are "singlet-like". Remember from introductory chemistry that transition metal ligands are names with a specific nomenclature. Some are shown in Figure $14$. Figure $14$: Common oxygen transition metal ligands In naming η is the hapticity (the number of atoms of a ligand attached to a metal) and µ is the number of metal atoms bridged by a ligand. For Fe ions and oxygen ligands, some examples include FeIII–O2 (superoxo) and FeIV=O (ferryl-oxo) If the aromatic substrate get oxidized, something must get reduced. That something is of course O2. Electrons for the reduction of O2 come from the oxidized substrate but also by injection of electrons through NAD(P)H. Cytochrome P450 are microsomal protein and most require another protein, NADPH-cytochrome P450 reductase (CPR). Before we look at the structure and mechanism of cytochrome P450, lets looks at this microsomal protein first. CPR catalyzes this reaction: NADPH + oxidized CytoP450 (Fe3+-heme b) + H+ ↔ NADP+ + Oxidized CytoP450 (Fe2+-heme b) The protein contains multiple domains and resulted from a fusion of gene from flavodoxin, which binds FMN) and a FAD reductases. In addition is contains an NADP binding domain. It ultimately accepts a hydride ion (2 electrons) from NADPH and then transfers electrons to FMN (in one electron steps). These are used to reduce the heme which then activates dioxygen (oxidation number of 0) for hydroxylation reactions by cytochrome P450. In the hydroxylated organic product and in water, oxygen has an oxidation number of -2. Figure $15$ below shows an interactive iCn3D model of rat NADPH-cytochrome P450 reductase (1AMO). Figure $15$: Rat NADPH-cytochrome P450 reductase (1AMO). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...nMh9ByRbsWk4B8 NADP+ is labeled as NAP. The enzyme is quite unique in that it has binding sites for both FAD and FMN (similar to nitric-oxide synthase). The flavins are aligned for electron transfer. The cleft in the enzyme presumably binds cytochrome P450. The protein has short (20 amino acid) lumenal sections, followed by 20 amino acid membrane-spanning helix (not shown in the above structure) followed a large cytosolic domain, where it can interact with cytochrome P450, which is found in multiple places including the ER, microsomal, and mitochondrial membranes as well as the cytoplasm.. The cytochromes P450s containing a heme, which instead of reversibly carrying dioxygen (as in myoglobin and hemoglobin), activates dioxygen for hydroxylation reactions involving aromatic, nonpolar substrates. Hydroxylation of these substrates increases their solubility which facilitates their elimination from the body. Epoxide intermediates or products are often produced, which can open up through nucleophilic attack using an alcohol (sugar derivative), as illustrated in Figure $16$. Figure $16$: Cytochrome P450 hydroxylation reaction to increase target solubility This type of reaction also converts steroids to different biologically active one. The hydroxylation reaction and can also lead to reactions with amines, including those on nucleotide bases in DNA, resulting in the formation of large adducts, as illustrated in Figure $17$. Figure $17$: Cytochrome P450-mediated formation of carcinogens. Hence cytochrome P450 can actually activate aromatic substrates to become carcinogens. The cytochrome P450s family of genes/proteins are inducible on exposure to nonpolar aromatic molecules such as dioxin. These nonpolar molecules can enter the cytoplasm where they bind to the arylhydrocarbon receptor (AhR) which is bound to a heat shock protein, Hsp90. Upon binding of dioxin, TCDD, for example, the AhR.TCCD complex dissociates from Hsp90, and migrates to the nucleus where it binds a protein called Amt. The AhR-Amt complex serves as an enhancer/transcription factor, facilitating the transcription of the cytochrome P450 genes. Figure $18$ illustrates the activation of cytochrome P450 gene expression on exposure to nonpolar aromatic molecules such as dioxin. Figure $18$: Activation of cytochrome P450 and other gene expression on exposure to nonpolar aromatic molecules such as dioxin On binding to a ligand, the AhR is activated and enters the nucleus, where it binds to ARNT on the aryl hydrocarbon response element (AhRE) and promotes transcription of downstream genes including cytochrome P450 family 1 subfamily A member 1 (CYP1A1) and interleukin-1 (IL-1). CYP1A1 is in the CYP1A family that promotes activation of procarcinogens a well as hydroxylation of steroid hormones like estrogens. It also participates in the metabolism of steroidal hormones including estrogens. ARNT is the aryl hydrocarbon receptor nuclear translocator, AhRE the aryl hydrocarbon response element; CYP1A1, XAP2 is the aryl hydrocarbon receptor interacting protein, AHRR is the aryl-hydrocarbon receptor repressor, IL-11 is interleukin 17, Hsp90 is heat shock protein 90 and p23 is prostaglandin E synthase 3 Dioxin has been shown to affect estrogen-mediated activities. Estrogens, small hydrophobic hormones derived from cholesterol, enter cell and bind to cytoplasmic estrogen receptors, which then dimerize and bind to the estrogen response element (ERE), initiating transcription. Tamoxifen, a drug derived from the yew plant, blocks the biological effects of the estrogen receptor. Although it binds to the estrogen receptor, it doesn't elicit the same conformational changes in the protein, which prevents the bound receptor from binding to the estrogen response element and recruiting other proteins needed for estrogen-dependent gene transcription. It is used in chemotherapy and prevention of estrogen-dependent breast cancer cells. How does dioxin interfere with estrogen signaling? Ahr and ARnt contain a basic helix-loop-helix motif which mediate their interaction with DNA. Upon binding of the complex, detoxification genes are activated. The dioxin-Ahr-Arnt complex can also bind to the estrogen receptor, which can then lead to activation of genes containing an estrogen response element (ERE) in the absence of estrogen. However, if estrogen is present, inhibition of gene expression from ERE is observed. Dioxins can be potent dysregulators of estrogen-induced gene expression. Such changes in estrogen activity could help to explain the pro- and inhibitory effects of dioxin on estrogen-mediated cellular responses and possible effects of dioxin on the immune system and on cancer development. Given the importance of the cytochrome P450s, we'll offer two variant portrayals of their mechanism. Figure $19$ shows the overall catalytic cycle of the enzyme with associated redox changes in the generic substrate, RH, and the Fe heme ion. Figure $19$: Catalytic cycle of cytochrome P450 with the generic substrate RH and the Fe heme ion The key hydroxylating agent appears to be formal FeO3+ shown in step 7. A possible mechanism for the hydroxylation of estrone by cytochrome P450 from Bacillus megaterium is shown below in Figure $20$. It follows the general catalytic cycle shown above. Estrone + NADH + H+ + O2 → 2-hydroxyestrone + NAD+ + H2O Figure $20$: Possible mechanism for the hydroxylation of estrone by cytochrome P450 from Bacillus megaterium after https://www.ebi.ac.uk/thornton-srv/m...csa/entry/699/ As with the previous generic mechanism, the two added electrons derive from NADH which passed a hydride to FAD which passes single electrons on to FMN which them passes them onto the heme. The exact species for several parts of the mechanism are not completely clear. For instance, a Fe4+-oxo complex has been proposed. Let's shown the structure of a human cytochrome P450 1A1 that is active in drug metabolism and also in the activation of benzo[a]pyrene, a component of cigarette smoke, into a carcinogen. Figure $21$ below shows an interactive iCn3D model of Human Cytochrome P450 1A1 in complex with alpha-naphthoflavone (4I8V). Figure $21$: Human Cytochrome P450 1A1 in complex with alpha-naphthoflavone (4I8V). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...JGtJeHG1qswN59 The shows in sticks and labeled the active site heme and an inhibitor, α-naphthoflavone (labeled BHF) bound and enclosed in the active site. It's 2-phenyl group points toward the heme Fe ion. Dioxygenases An example of a dioxygenase is the cyclooxygenase activity of prostaglandin synthase. This enzyme, often just called cyclooxygenase or COX, is an integral membrane protein found in the ER membrane, and is a homodimer (with two hemes). It catalyzes two different reactions. One is the addition of two dioxygens to arachidonic acid - 20:4Δ5, 8, 11, 15 (which is liberated from the C2 position of phospholipid membranes by phospholipase A2 upon appropriate signaling) to form prostaglandin PGG2. This molecule, with 5 chiral centers, arises from arachidonic acid, which has one. The cyclooxygenase activity is buried in the membrane, from where the arachidonic acid can readily access its active site. The active site is at the end of a hydrophobic channel (arachidonic acid binding site) and stretches from the membrane-binding region to a buried heme. PGG2 can be further metabolized to PGH2 by the addition of two electrons to PGG2 by the hydroperoxidase activity of the enzyme, located at the other end of the enzyme. This activity forms an alcohol from the peroxide functional group in PGG2. There is one heme per monomer, which acts in both the cyclooxygenase and peroxidase activities. Each monomer of the dimer has both enzymatic activities. A possible abbreviated reaction mechanism is shown in Figure $22$. Figure $22$: A possible abbreviated reaction mechanism for dioxygenase cyclooxygenase. In summary: • The carboxylate of arachidonic acid is coordinated to Arg 120 and Tyr 355. • The C13 pro(S) H atom of arachidonic acid is close to Tyr 385 which allows its abstraction. • This results in a radical centered on C11 that reacts with dioxygen to form a peroxyl radical • Attack by dioxygen at C11 occurs from the side of the substrate opposite to that of hydrogen abstraction • The oxygen radical at C11 cyclizes by attacking C9. The C13 proS hydrogen atom (not proton) is remove from bound arachidonic acid by a free radical form of Tyr 385, which acts as an oxidizing agent. A site-specific mutants in which Tyr 385 is replaced by Phe is inactive. How is the Tyr free radical formed? Based on single electron standard reduction potential (0.9 V for Tyr and -0.2 to + 0.2 V for Fe3+ in the bound heme), it appears that the heme iron is not a potent enough oxidizing agent to accomplish this task. However, oxygen bound to the heme iron could be converted to a peroxide and form an Fe4+-oxo complex (which has also been proposed for cytochrome P450). The Fe4+ ion is a more potent oxidizing agent (standard reduction potential of approximately 1 V, sufficient for oxidation of Tyr 385. Another possibility is that the peroxide activator (in the formation of the ferryl-oxo ligand) is NO (nitric oxide, a free radical). NO is formed by immune cells (like macrophages) on immune activation. The NO might react with superoxide (also a radical, possibly formed during an oxidative burst in macrophages during immune stimulation) to form peroxynitrite (NO3-). This can donate an oxo group to the Fe3+ to form the Fe4+-oxo complex, which could then oxidize Tyr to the free radical form. There might be other mechanisms as well to generate the Tyr free radical, since just adding organic peroxides to the enzyme will generate it.. After abstraction of the proS H atoms, a carbon-centered free radical at C11 results which reacts with oxygen as shown below. The exact form of oxygen that reacts is unclear, but presumably is either a peroxy or activated singlet form. Oxidases This class of enzymes does not incorporate dioxygen into an organic substrate. Rather it accepts electrons released from an organic substrate, through intermediate electron carriers (such as ubiquinone and cytochrome C) to form superoxide (as in NADPH-oxidase), hydrogen peroxide (as in xanthine oxidase) or water (as in cytochrome C oxidase). The mechanism of cytochrome C oxidase again supports our expectations about enzymes that use dioxygen. Dioxygen binds metals in the enzyme. One oxygen atom binds a heme Fe2+ of cytochrome a3 which is bound to the enzyme, while the other binds a Cu1+ of Cu B. All oxygen reduction intermediates remain bound to the enzyme. Four electrons are added from four different cytochrome C molecules, which serve as mobile carriers of electrons. We will explore cytochrome C oxidase in great detail in Chapter 19.1: Electron-Transfer Reactions in Mitochondria. Figure $23$ shows cartoon versions of several oxidases. Figure $23$: Examples of oxidases Another example of an oxidase is monoamine oxidase. Mitochondrial monoamine oxidase catalyzes the oxidative deamination of certain neurotransmitters after they have been taken up by post-synaptic neurons, in a process of inactivation. The reaction is shown in Figure $24$. Figure $24$: Monoamine oxidase reaction A Schiff base is formed which is then hydrolyzed, incorporating unlabeled oxygen into the oxidized molecule. Biological Oxidations: Methane to CO2 In the previous chapter section, we discussed the progressive oxidation of methane by 2 electron loses to form methanol, formaldehyde, formic acid and CO2, with a progressive increase in oxidation number for the C by +2 (from -4 in methane to +4 in CO2), as reviewed in Figure $25$. Figure $25$:: Progressive stages in the oxidation of methane Methanotrophs are aerobic bacteria that use methane as a source of energy, converting it in a series of two electron oxidations as shown above, to carbon dioxide. The enzymes involved in this sequential process are methane monooxygenase, methanol dehydrogenase, formaldehyde dehydrogenase, and formate dehydrogenase. Methane monooxygenase exists in a soluble and membrane form, both of which are part of a larger complex. Both have a hydroxylase (which uses dioxygen to add O to methane) and the membrane form has recently been shown to be associated with have methanol dehydrogenase in a larger complex consisting of trimers of each enzyme (the hydroxylase and the dehydrogenase). Heme Proteins So far in this course, we have examined three different kinds of heme proteins.. • The first, hemoglobin (and myoglobin) serve as carriers of dioxygen. Even though they bind one of the best oxidizing agents around (dioxygen), the heme Fe2+ does not get oxidized to Fe3+. If it does, as in the case of met-Hb, the protein looses it ability to carry oxygen. • Cytochrome C, on the other hand, does not bind dioxygen but rather serves as a carrier of electrons which get passed to dioxygen in Cytochrome C oxidase. Its Fe ion readily cycles between the 2+ and 3+ states as it serves as an electron carrier. • Finally, the Fe2+ in the heme of the cytochrome P450s (so named since they have an absorbance maximum at 450 nm when they bind CO) does both. It binds dioxygen and cycles between the 2+ and 3+ states as it activates dioxygen for hydroxylation reactions. The structure of the heme and amino acid ligands, along with their absorbance spectra, are shown in Table $3$ below. hemoglobin cytochrome C Cytochrome P450 P450: Fujishiro et al. J Biol Chem. 2011 Aug 26; 286(34): 29941–29950. Published online 2011 Jun 30. doi: 10.1074/jbc.M111.245225. CC BY license. Cyto C: Hulko et al. December 2011. Sensors 11(6):5968-80. DOI:10.3390/s110605968. Creative Commons Attribution 3.0 Unported Hemoglobin: Nitzan et al. July 2014, Medical Devices: Evidence and Research 7(1):231-9. DOI:10.2147/MDER.S47319. Creative Commons Attribution-NonCommercial 3.0 Unported How could heme serve such diverse functions? We can explain this by referring to one of the main themes of the course - structure mediates function. The environment of each heme must be different. Clearly the protein ligands coordinating the Fe ions are different. The 5th ligand is the proximal His in hemoglobin while dioxygen binds to the 6th site. In cytochrome C, the 5th and 6th ligands are His and Met, respectively. In cytochrome P450, the 5th site is occupied by Cys, and the 6th by dioxygen. Presumably the environments surrounding the hemes are different as well. Once again, we have seen analogous example in which chemical properties are influenced by the microenvironment. The pKa of a given amino acid side chain can vary considerably depending on the polarity of the local environment. Likewise, the standard reduction potential of tightly bound FAD/FADH2 depends on the microenvironment. As we have seen (from the study of heme proteins and the oxidative enzymes of cells), transition metals such as Fe, Zn, and Cu have vital biological roles as binding sites and cofactors in many reactions. Yet they also pose problems since they can lead to oxidative damage in cells. As we saw with cytoplasmic metallothioneins, which bind to heavy metals and protect the cell from such damage, many proteins are involved in binding and regulation of transition metals in the cell. Integral membrane proteins are required to bind and transport these cations into the cytoplasm. Other proteins act as sensors of transition ion concentration (such as latent transcription factors which bind heavy metals and become active transcription factors for metallothioneins. Others act as chaperone proteins which bind metal ions and transfers them to apometalloproteins. Recent work has suggested transporters and chaperones involved in metal ion biology bind these ions with unusual coordination geometry, which presumably facilitates transfer of the ion to the apo-target protein. The transition metals Zn and Fe are often found in E. Coli at a concentration of 0.1 mM, compared to Cu and Mn which are present at concentrations from 10 to 100 μM. Also, about one third of all proteins demonstrate specific binding of metal ions and can be classified as metalloproteins. Mass balance suggests that metal ions would be distributed in proteins with low, intermediate, and high metal binding affinity as well as in free pools, which which potentially be toxic to cells. Metalloproteins, depending on their Kd for metal ion binding, would hence be in various state of ligation. The free concentration of some ions (Cu and Zn) is so low that newly synthesized apoproteins which bind these ions would not obtain the ion from the free pool. In such cases, metal chaperones would be required.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/12%3A_Bioenergetics_and_Biochemical_Reaction_Types/12.04%3A_Biological_Oxidation-Reduction_Reactions.txt

• 13.1: Glycolysis In this section, we will provide you with a historical overview of glycolysis and introduce you to the 10 enzymatic reactions in the pathway. Our main goal is to understand how the oxidation of our major food molecules, sugars in the case of glycolysis, can lead to ATP synthesis. • 13.2: Fates of Pyruvate under Anaerobic Conditions- Fermentation Fermentation is a process in which a fuel molecule is broken down in an anaerobically (or without oxygen), to produce energy. One of the most notable pathways that can utilize fermentation is glycolysis, which we have just described. During glycolysis, glucose is converted to pyruvate, yielding a total of 2 ATP energy rich molecules. This occurs in anoxic conditions (or without the requirement of oxygen). • 13.3: Gluconeogenesis Gluconeogenesis is a metabolic pathway that results in the generation of glucose from non-carbohydrate carbon substrates such as lactate, glycerol, and glucogenic amino acids. It is one of the two main mechanisms humans and many other animals use to keep blood glucose levels from dropping too low (hypoglycemia). The other means of maintaining blood glucose levels is through the degradation of glycogen (glycogenolysis). • 13.4: Pentose Phosphate Pathway of Glucose Oxidation The pentose phosphate pathway (PPP), also known as the pentose phosphate shunt, is an important part of glucose metabolism. The PPP branches after the first step of glycolysis and consumes the intermediate glucose 6-phosphate (G6P) to generate fructose 6-phosphate (F6P) and glyceraldehyde 3-phosphate (G3P) through the oxidative and non-oxidative branches of the PPP. Thumbnail: Due to its role in the pentose phosphate pathway and the Calvin cycle, D-Ribose-5-phosphate isomerase is highly conserved in most organisms, such as bacteria, plants, and animals. It plays an essential role in the metabolism of plants and animals, as it is involved in the Calvin cycle which takes place in plants, and the pentose phosphate pathway which takes place in plants as well as animals. (CC BY-SA 4.0; Ishikawa, K., Matsui, I., Payan, F., Cambillau, C., Ishida, H. et al. via Wikipedia) 13: Glycolysis Gluconeogenesis and the Pentose Phosphate Pathway Search Fundamentals of Biochemistry Introduction In this chapter, we will provide you with a historical overview of glycolysis and introduce you to the 10 enzymatic reactions in the pathway. Our main goal is to understand how the oxidation of our major food molecules, sugars in the case of glycolysis, can lead to ATP synthesis. Before we begin our journey into the glycolytic pathway, it is useful to review the concept of free energy within reactions. For a reaction to be spontaneous, the change in the free energy within the system must be negative. From the equation in Figure $1$ you can see that the change in free energy of a reaction is dependent on the concentration of the reactants and the products, as well as the temperature within the system. You will note that some reactions of glycolysis and other metabolic pathways that we will investigate are not favored. Thus, it is necessary to drive the reaction to become spontaneous by either coupling the reaction with a spontaneous reaction that can generate enough free energy to drive the nonspontaneous reaction forward, or by using Le Chatelier’s principle and removing the products from the enzyme’s area as soon as they are made. This will help drive the reaction in the forward direction as it will reestablish equilibrium. In this way, a small amount of product can be formed spontaneously. Essentially, the continued removal of the product or the addition of excess reactant will drive the reaction forward. It is of note that the temperature of the reaction can also influence the change in free energy. However, in biological systems, temperature changes that will significantly affect the change in free energy usually are not compatible with maintaining most life forms. Thus, it will not be a large consideration in the context of our metabolic discussions, here. When reactions are coupled together to obtain a spontaneous reaction, the overall free energy change for a chemically coupled series of reactions is equal to the sum of the free energy changes of the individual steps, as noted in Figure $2$. The metabolic reactions of carbohydrates and other food molecules play an important role in generating energy within living systems in the form of ATP. For carbohydrates, this begins with the metabolic process known as glycolysis (or the breakdown, "lysis", of sugars, "glyco") At the end of the 1910s Otto Meyerhof mapped some of these metabolic conversions by measuring heat trends and oxygen consumption in frog muscles. When the muscle is working, lactic acid is formed from carbohydrates, and Otto Meyerhof showed that during recovery, this is followed partly by the burning of lactic acid and partly by reprocessing of lactic acid to carbohydrates. Concurrently, Archibald Hill was also outlining these processes in the muscles of frogs. In opposition to the prevailing view that mechanical movement and chemical processes were parallel sequences, Hill was able to show through measurements of heat generated by the mechanical processes that these were delayed compared to the movements. The chemical sequence consists of a work phase, which is not dependent on oxygen supply, and a recovery phase when oxygen is required. Together, their work has opened the door to understanding aerobic and anaerobic metabolism beginning with the process of glycolysis. They both shared the Nobel Prize in Physiology and Medicine in 1922 for their work in these processes Figure $3$. The glycolytic pathway consists of 10 enzymatic steps that convert glucose to pyruvate. This conversion generates a small amount of energy. The pyruvate can then be converted to lactic acid (lactate) in vertebrates or to ethanol in yeast in an anaerobic (or oxygen-independent) pathway or it can be fully oxidized to carbon dioxide in an aerobic (or oxygen-requiring) pathway that takes place within the mitochondria (which is shown in green in Figure $4$: ). Aerobic oxidation yields about 18 times as much energy as the anaerobic pathways causing them to be favored over anaerobic pathways.  Most animal tissues can only survive short anaerobic bursts that occur in isolation and don’t involve the entire organism. The aerobic pathway is required to sustain life. Yeast also prefers to grow using the aerobic, mitochondrial pathway. However, if oxygen is unavailable, yeast and other fungi can switch to anaerobic growth and produce ethanol as a byproduct. The production of alcoholic beverages through this fermentation process is quite popular. Figure $5$ provides a summary of the glycolytic pathway coupled to the oxidative phosphorylation pathway that occurs within the mitochondria. Figure $5$: A Summary of the Glycolytic and Oxidative Phosphorylation Pathways. The glycolytic pathway is shown on the left-hand side in blue. During aerobic metabolism, pyruvate would be converted to acetyl-CoA which then enters the Kreb cycle within the matrix of the mitochondria. Future chapters will focus on the reactions inside the mitochondria. Our focus in this chapter will be on the cytosolic reactions of the glycolytic pathway. The major reactions of glycolysis are shown in Figure $6$. The pathway can be broken down into two major sections: (1) the energy-consuming reactions and the (2) the energy-generating reactions. An adage states that ‘It takes money to make money’. The same can be thought about the glycolytic pathway. The first section requires an investment of energy to generate energy in the second half of the reaction pathway. In this pathway, glucose, a 6-carbon hexose, is converted to two, 3C molecules - pyruvate. Note that Figure $6$ shows the entire pathway using Lewis wedge/dash representations plus the anaerobic conversion of pyruvate to lactate. Figure $6$: Glycolytic pathway. The chemical steps of the glycolytic pathway are shown using Lewis wedge/dash representations. Enzymes required at each step are labeled in red. The energy-consuming steps encompass reactions 1 - 3, whereas the energy-producing steps occur in the second half of the pathway from reactions 4 - 10. The conversion of pyruvate to lactate by lactate dehydrogenase represents anaerobic respiration as it occurs within mammalian species. Glycolysis is the key anaerobic pathway for energy products in all organisms, except in lithotropes that use the oxidation of inorganic molecules for energy production. In aerobic systems, glycolysis provides the release of fast energy within the body as glycogen metabolism can quickly release free glucose for utilization. Given the centrality of glycolysis to all of life, we will explore each of the reactions in detail below. Reaction 1: Glucose → Glucose-6-Phosphate. ΔGo=-4.0 kcal/mol (-16.7 kJ/mol). The first step in glycolysis is catalyzed by enzymes known as hexokinases. The hexokinase family enzymes typically have broad specificity for various hexoses and catalyze the phosphorylation of carbon 6. Hexokinase phosphorylates glucose using ATP as the source of the phosphate, producing glucose-6-phosphate, a more reactive form of glucose. Notably, this reaction prevents the phosphorylated glucose molecule from continuing to interact with the GLUT transport proteins that can shuttle glucose into and out of the cell. Thus, once glucose is phosphorylated it can no longer leave the cell. Do note that both of these sugar forms (the free sugar and the phosphorylated version) can shift back and forth between the ring-closed and ring-opened conformation. Hexokinases require glucose to be in the closed conformation for the phosphorylation reaction. Figure $7$ shows the reaction which is catalyzed by hexokinase (a kinase that transfers the γ-phosphate from ATP to a hexose in the closed-ring form). Figure $7$ : Summary reaction - hexokinase This reaction is a nucleophilic substitution reaction on the gamma phosphate of ATP. A phosphoanhydride bond is broken in ATP as a phosphoester bond is made producing glucose 6-phosphate. Hence the reaction proceeds with a negative ΔGo. Vertebrates have 4 different types of hexokinases, I-IV (also called A-D). The regulation of these enzymes is presented in more detail in Chapter 15.5. Hexokinases I-III bind glucose more tightly as reflected by low KM values. Type IV or glucokinase binds it less tightly, and is found in high concentrations in vertebrate livers. Yeast has 3 isozymes, P1, PII or hexokinase B, and glucokinase. Most prokaryotic hexokinases are in the glucokinase (Type IV) class. In addition to hexokinases, there are also many other different types of sugar kinase enzymes. Notably, all sugar kinases must prevent the spurious hydrolysis of ATP by water, which can also be considered a phosphotransfer to water, instead of the preferred transfer of the γ phosphate of ATP to a sugar ROH (alcohol), an "alcoholysis" reaction. Hexokinase does this through an "induced fit" mechanism, in which glucose binding triggers a large conformation change to close off the active site to water. This conformational change is illustrated in Figure $8$ below which shows an interactive iCn3D model of the yeast hexokinase PI in the absence (2YHX) and presence (3B8A) of glucose. Figure $8$: Yeast hexokinase PI in the absence (2YHX) and presence (3B8A) of glucose. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...84YGEd1ob3th7A Within Figure $8$, the gray structure is hexokinase without glucose (2YHX). However, it has a glucose analog bound, o-tolyoylglucosamine (spacefill, yellow highlights), which binds at the glucose binding site but does not cause a subsequent conformation change. The cyan structure (in the actual iCn3D model) show the enzyme with glucose shown as colored sticks. Note the large conformational change on the actual binding of glucose, a classical example of an "induced fit" mechanism. Figure $9$ shows the surface of both enzymes using the same color coding as in the above figure These images better show how the active site of hexokinase is occluded in the glucose-bound form. This prevents water access and hydrolysis of bound ATP instead of "alcoholysis" of ATP (transfer of phosphate to glucose). The gray structure on left has the glucose analog bound which doesn't alter the global conformation of the protein. Figure $9$: Surface of human hexokinase I with bound o-tolyoylglucosamine (spacefill, left, 2YHX) and with bound glucose (spacefill, right, 3B8A) Hexokinase I is the key form in the brain. It forms a complex with porin in the mitochondrial outer membrane and ATP/ADP translocase or carrier protein in the mitochondrial inner membrane which facilitates the hexokinase reaction. The mechanism for the reaction of human hexokinase I is shown in Figure $10$ below. Figure $10$: Mechanism of human hexokinase I. Ribeiro AJM et al. (2017), Nucleic Acids Res, 46, D618-D623. Mechanism and Catalytic Site Atlas (M-CSA): a database of enzyme reaction mechanisms and active sites. DOI:10.1093/nar/gkx1012. PMID:29106569. https://www.ebi.ac.uk/thornton-srv/m-csa/entry/696/. Creative Commons Attribution 4.0 International (CC BY 4.0) License. Figure $11$ below shows an interactive iCn3D model of human hexokinase I with glucose and ADP in the active site (1dgk). Figure $11$: Human hexokinase I with glucose and ADP in the active site (1dgk). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...aQy4gf6A2bWtp8 The key catalytic residues are shown in colored sticks and labeled. Glucose, ADP, and PO42- are shown as colored spheres. Note that hexokinase I, as well as forms II and III, have spatially distinct halves similar to those in the yeast enzymes. Forms I-III have a molecule weight of about 100K. Form IV (glucokinase) has a molecular weight of 50K and is similar to the distinct halves of I-III. suggesting that I-III arose through gene duplication. Each half also binds glucose and ADP but the N-terminals of I and III are catalytically inactive. Hexokinase II, with both halves active, is the main enzyme involved in glycolysis in most mammalian tissues. Other differences are that glucose-6-phosphate, a product, inhibits I-III. PO42- relieves G6P product inhibition in form I but not the others. ADP binding at multiple sites in hexokinase I likely causes a conformational change that affects structure and activity. Sugar kinases in general We will see many kinases that use ATP to phosphorylate sugars, so it is useful to explore both their differences and similarities at the beginning of our studies on carbohydrate metabolism. Figure $12$ below shows the comparative structures of the five carbohydrate kinase classes. Figure $12$: Structures of the five carbohydrate kinase classes. All images are shown in rainbow format (blue: N-terminus, red: C-terminus). Roy, S.; Vivoli Vega, M.; Harmer, N.J. Carbohydrate Kinases: A Conserved Mechanism Across Differing Folds. Catalysts 2019, 9, 29. https://doi.org/10.3390/catal9010029. Creative Commons Attribution License Panel (a) shows the structure of human glucokinase (hexokinase class; PDB (protein data bank) ID: 4IWV). Panel (b) shows the structure of Bacillus subtilis fructokinase dimer: the second molecule shown in raspberry (ROK kinase class; PDB ID: 1XC3 [17]). Panel (c) shows the structure of Escherichia coli ribokinase (ribokinase class; PDB ID: 1RKD; [18]). Panel (d) shows the structure of Aquifex aeolicus IspE (GHMP kinase class; PDB ID: 2V2Z; [19]). Panel (e) shows the structure of human PIK3C3 (phosphatidylinositol phosphate kinase class; PDB ID: 3IHY). Table $1$: Overview of the Five Carbohydrate Kinase Classes. Carbohydrate Kinase Family Common Substrates Native Phosphate Donors (Minor Donors in Parentheses) Pfam ID Hexokinase Glucose, mannose, fructose ATP (ITP) PF00349, PF03727, PF02685 ROK Kinase Glucose, allose, fructose, N-acetylglucosamine, N-acetylmannosamine ATP (polyphosphate) PF00480 Ribokinase Ribose, 2-deoxy-d-ribose, adenosine ATP, ADP (GTP, ribonucleotide) PF00294 GHMP Kinase Galactose, N-acetylgalactosamine, ATP (GTP, ITP) PF00288 Phosphatidylinositol kinase Phosphatidylinositol, phosphatidylinositol phosphates ATP (GTP) PF00454, PF01504 ROK is a bacterial Repressor, Open reading frame, Kinases are predominantly bacterial enzymes. Ribokinases include adenosine kinases, fructokinases, and phosphofructokinases. GHMP Kinases include Galactokinase, Homoserine kinase, Mevalonate kinase, and Phosphomevalonate kinase. Roy et al. Ibid Many of these kinases are regulated by the binding of allosteric effectors. Figure $13$ shows a common mechanism for all, using glucose as an example substrate. Figure $13$: Generic reaction mechanism for carbohydrate kinases (hexokinase shown). Roy et al, Ibid. The generic reaction is initiated by a catalytic base abstracting a proton from the reactive hydroxyl (left). The oxygen atom then attacks the -phosphate of ATP (second left), forming a pentacoordinate transition state (second right). This is stabilized by a divalent cation, and by the protein (not shown). This transition state resolves, leaving ADP and the phosphorylated carbohydrate (right). Reaction 2: Glucose-6-Phosphate ↔ Fructose-6-Phosphate. ΔGo=+0.4 kcal/mol (+1.7 kJ/mol) In the second step of glycolysis, the phosphoglucose isomerase (PGI) converts glucose-6-phosphate into one of its isomers, fructose-6-phosphate. Recall that an isomerase is an enzyme that catalyzes the conversion of a molecule into one of its isomers. In this reaction, the aldose, glucose-6-phosphate, is converted to the ketose, fructose-6-phosphate. This conversion is essential to allow the eventual split of the sugar into two three-carbon molecules. We'll present this reaction, catalyzed by phosphoglucose isomerase (PGI), in several different types of representations (chair, wedge/dash, and Fischer projection), as shown in Figure $14$ below. Figure $14$: Summary reaction, phosphoglucoisomerase From a functional perspective, seen more clearly from the linear Fischer structure, it is evident that the C=O has been moved to the C2 position to create the ketose structure. The carbonyl O is now positioned to be an electron sink facilitating electron flow for reaction 4. This isomerization reaction would be expected to have a ΔGo of about 0. It has been proposed that the mechanism of PGI requires the enzyme to open the Glucose-6-Phosphate ring before the actual isomerization step, which proceeds through the formation of a cis-enediol intermediate and keto-enol tautomers before conversion back to the cyclic form. Figure $15$ below shows a proposed mechanism for rabbit phosphoglucose isomerase Figure $15$: Mechanism for phosphoglucose isomerase. Ribeiro AJM et al. Ibid Figure $16$ below shows an interactive iCn3D model of rabbit phosphoglucose isomerase with bound 6-phosphogluconic acid (1DQR) Figure $16$: Rabbit phosphoglucose isomerase with bound 6-phosphogluconic acid (1DQR). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...xo1SgF1ZpAySm9 One monomer of the dimer is shown in gray and the other in cyan. The catalytic residues Lys518 and His388 are shown in both subunits in colored sticks and labeled. Additional residues that contribute to specificity (Ser209, Ser159, Thr214, Thr217, and Thr211) are shown in the gray subunit with color sticks and labeled. 6-phosphogluconic acid, a competitive inhibitor, is shown in color spacefill. The enzyme also has other functions in addition to its role in glycolysis, so it is a member of a group called "moonlighting" (a term that refers to working at a secondary job). proteins. Outside of the cell, phosphoglucoisomerase acts as a nerve growth factor and cytokine. It is also called autocrine motility factor (PGI/AMF) and its cytokine activity is associated with aggressive cancers. Reaction 3: Fructose-6-Phosphate → Fructose-1,6-Bisphosphate. ΔGo=-3.4 kcal/mol (-14 kJ/mol) The third step is the phosphorylation of fructose-6-phosphate, catalyzed by the enzyme phosphofructokinase-1 (PFK1). A second ATP molecule donates a high-energy phosphate to fructose-6-phosphate, producing fructose-1,6-bisphosphate. The term bisphosphate is used when two phosphate groups are joined to a molecule at different positions on the molecule. In this case, one phosphate is at the 1-carbon position and the other is at the 6-carbon position. This differs from the term diphosphate, which is used when the phosphate groups are joined in a sequence, as in the molecule ADP. In ADP, both phosphate groups are joined in tandem from the 5-carbon position on the ribose ring structure. In the glycolytic pathway, PFK1 is a rate-limiting enzyme. The mechanism of PFK1 regulation is discussed in greater detail in Chapter 15.5.  However, we will introduce the process here. Essentially, the enzyme is sensitive to the energy load within the cell. Recall that ATP is a recycled molecule and exists in a pool of interconverting  ATP, ADP, and AMP. A chief outcome of glycolysis is to shift the pool towards increased levels of ATP to drive endergonic processes like muscle contraction.  PFK1 activity is sensitive to the ATP:ADP ratio within the cell. PFK1 is more active when the concentration of ADP is high and the concentration of ATP is low, and it is conversely less active when ADP levels are low and the concentration of ATP is high. This is a type of end-product inhibition since ATP is the end product of glucose catabolism. Note, however, that ATP is also a substrate for PFK1. ATP serves as the phosphate donor in the reaction and is required in the process. This is the second energy-intensive step in the glycolytic pathway. At the end of the PFK1 step, a total of 2 ATP molecules have been broken down in the glycolytic pathway. The PFK1 enzymatic step is also important within the glycolytic pathway as it is the committed step in the pathway. Glucose-6-phosphate may be used for other purposes within the cell, and the isomerase step that converts glucose-6-phosphate to fructose-6-phosphate is readily reversible. The PFK1 enzyme only works in the forward direction to create fructose 1,6-bisphosphate. It cannot in the reverse direction and recover the reactant. Thus, it is thought of as the committed step in the glycolytic pathway as the fructose-1,6-bisphosphate will predominantly be converted into pyruvate through the remaining enzymatic steps. The reaction catalyzed by PFK1 is shown in Figure $17$ below. Figure $17$: Summary reaction - phosphofructokinase By phosphorylating this intermediate, both products of the cleavage of this 6C molecule will be phosphorylated, keeping both more readily inside the cell. This reaction is a nucleophilic substitution reaction on the gamma phosphate of ATP. A phosphoanhydride bond is broken in ATP as a phosphoester bond is made producing fructose-1,6-bisphosphate. As in reaction 1, this reaction proceeds with a negative ΔGo The PFK1 enzyme exists as a tetramer and as with another "famous" tetramer, hemoglobin, it can exist in T and R symmetry states. Hence it is an allosteric enzyme and has many allosteric regulators. One important allosteric activator of eukaryotic (but not prokaryotic) PFK is fructose-2,6-bisphosphate. This is formed by a separate phosphofructokinase enzyme named PFK2. This regulatory pathway will be described in greater detail in Chapter 15.5. Hence the number 1 is added to the name of the glycolytic enzyme, PFK1, that forms fructose 1,6-bisphosphate, to indicate the position of phosphorylation. Mammals have three isoforms of PFK1, muscle (PFKM), liver (PFKL), and platelet (PFKP). The mechanism for E. Coli phosphofructokinase is shown in Figure $18$ below. A metal cofactor (Mg2+) coordinates the positioning of the ATP and stabilizes the gamma phosphate for nucleophilic attack by the fructose alcohol group at position-1. Activation of the fructose alcohol group is mediated by proton abstraction with a coordinated Asp127 of PFK1. Figure $18$: Proposed mechanism for E. Coli phosphofructokinase. Ribeiro AJM et al. Ibid Figure $19$ below shows an interactive iCn3D model of the E. Coli phosphofructokinase with bound F1,6-bisphosphate and ADP products (1PFK). (long load) Figure $19$: E. Coli phosphofructokinase with bound F1,6-bisphosphate and ADP products (1PFK). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...t4KKgLAGzHrzo8 Each monomer of the tetramer is shown in a different color. The gray monomers show key binding and catalytic residues described in the mechanism above. F1,6-bisphosphate is shown as spacefill and ADP is shown as sticks. The structure of the human platelet PFK1 tetramer has been determined in the presence of ATP and ADP. Figure $20$ below shows an interactive iCn3D model of the human phosphofructokinase-1 dimer (for clarity) in complex with ATP and Mg (4XYJ). (long load) Figure $20$: Human phosphofructokinase-1 dimer in complex with ATP and Mg (4XYJ). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...AKfonyGsNq6Cs9 (long load). Here is a link to the ADP bound structure: https://structure.ncbi.nlm.nih.gov/i...N81MBS4PZfSQp9 The monomers are shown as colored surfaces with secondary structures underneath. Note that side chains from each monomer in the dimer contribute to binding and catalysis. The actual PDB structure is tetrameric. The structures have an E173S mutation. Figure $21$ shows the differences in orientation of the key side chains in the ADP structure (left) and the ATP structure (right) that are key in the conformation changes in the enzyme on binding substrate. ADP bound form of human PFK (4XKJ) ADP bound form of human PFK (4XYJ) Figure $21$: Conformation changes in the active site of PFK on ATP hydrolysis Reaction 4: Fructose-1,6-bisphosphate → Dihydroxyacetone phosphate (DHAP) + Glyceraldehyde 3-phosphate (G3P). ΔGo= + 5.7 kcal/mol (+24 kJ/mol) The newly added high-energy phosphates further destabilize fructose-1,6-bisphosphate. The fourth step in glycolysis employs an enzyme, aldolase, to cleave fructose-1,6-bisphosphate into two three-carbon isomers: dihydroxyacetone phosphate and glyceraldehyde-3-phosphate. This reaction is shown in Figure $22$ below in both wedge dash and Fischer projections. Figure $22$: Summary reaction - aldolase This is the first C-C bond cleavage within glucose on the path to complete cleavage during aerobic respiration and release of 6 carbon dioxide molecules per glucose metabolized. This reaction is the reverse of an aldol condensation when an enol or enolate reacts with a carbonyl C to form an adduct. Note that both products are phosphorylated. The reaction is not thermodynamically favored but is pulled in the forward direction by the utilization of the product in subsequent reactions in the pathway. Table $2$ Characteristics of the Three Classes of Aldolases (I, IA, and II) and the Organisms in Which They are Found. Pirovich et al, Frontiers in Molecular Biosciences, 8 (2021), https://www.frontiersin.org/article/...lb.2021.719678. AUTHOR=Pirovich David B., Da’dara Akram A., Skelly Patrick J. Creative Commons Attribution License (CC BY). Class I Aldolase These enzymes proceed through a Schiff base intermediate between a reactive lysine and the reactant/product. The enzyme is favored in the reverse direction and it's perhaps easier to see the mechanism presented in that fashion. In rabbits, the muscle Class I aldolase (RAMA) uses Lys-229 to form a Schiff base with DHAP as shown in a mechanism presented in Figure $23$ below. Figure $23$: Class I aldolases - general mechanism (after Bolt et al., Arch Biochem Biophys. 2008 June 15; 474(2): 318–330) Only the pro(S) proton of the dihydroxyacetone phosphate C3 carbon is removed and effectively exchanged with the glyceraldehyde-3-phosphate substrate. Figure $24$ below reviews how the pro(R) and pro(S) hydrogens can be visually differentiated by replacing one with a deuterium and determining the stereochemistry of the now chiral C3. Figure $24$: Visualization of the Pro(R) and Pro(S) Hydrogens on DHAP Figure $25$ shows key active site residues in the active site of a Class I aldolase from rabbit muscle. The first step in Schiff base formation with Lys229 is shown. Figure $25$: Catalytic residues in rabbit muscle Class I enolase. Figure $26$ shows an interactive iCn3D model of the dihydroxyacetone phosphate enamine intermediate in fructose-1,6-bisphosphate aldolase from rabbit muscle (2QUT) Figure $26$: Dihydroxyacetone phosphate enamine intermediate in fructose-1,6-bisphosphate aldolase from rabbit muscle (2QUT). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...uR5KZBsfN8QQh9 Only one monomer of the four in the homotetramer is shown. The ligand, 1,3-dihydroxyacetonephosphate, is shown in spacefill with CPK colors. It is Schiff base linkage with Lys229. The key catalytic residues are shown and labeled. The reaction proceeds between eneamine and iminium covalent intermediates. A K146M mutation (in the active site) decreases enzyme activity and allows the trapping of the K229 eneamine intermediate. A key tyrosine (Y363) in its deprotonated state formed in the presence of the iminium phosphate and a local water appears to abstract the C3 pro(S) proton to form the enamine. Class II Aldolase Figure $27$: Figure $27$: Class II aldolases - general mechanism (after Bolt et al, ibid) Reaction 5: DHAP ↔ G3P. ΔGo= +1.8 kcal/mol (+7.5 kJ/mol) This reaction, catalyzed by triose phosphate isomerase (TPI), is shown in Figure $28$. Figure $28\: Summary reaction - triose phosphate isomerase. This is another simple isomerization reaction. Only one product, glyceraldehyde-3P continues on in glycolysis, so only one enzyme is needed to metabolize the cleavage products of this reaction further. As in other isomerization reactions, the ΔGo is close to 0. A proposed mechanism for this reaction is shown in Figure \(29 below. Figure \(29$: Proposed mechanism for triose phosphate isomerase (after Bolt et al, ibid) Note that the reaction employees the deprotonation of a neutral imidazole side chain to form an imidazolate anion. This would not likely be favorable, given its pKa value. The are four possible conserved proton donors with more reasonable pKas in the active sites of TPIs, including K12, H95, E97, and E165. Mutations of E97 to E97Q and E97D lead to a 4000-fold reduction in kcat (E97Q) but only a 100-fold reduction for E97D suggesting tha30}\) shows an interactive iCn3D model of the chicken triosephosphate isomerase-phosphoglycolohydroxamate complex (1TPH) Figure $30$: Chicken triosephosphate isomerase-phosphoglycolohydroxamate complex (1TPH). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...AFXpt9kg5aki5A The four key conserved residues are shown in the gray monomer of the homodimer. Reaction 6: G3P ↔1,3 BPG. ΔGo= +1.5 kcal/mol (+6.1 kJ/mol) This reaction, catalyzed by glyceraldehyde-3-phosphate dehydrogenase, is shown in Figure $31$ Figure $31\: Summary reaction - glyceraldehyde-3-phosphate dehydrogenase This is a big reaction! Note that the ΔGo is close to 0 but look at what happened. The carbonyl O in G3P has been oxidized to the form of a mixed anhydride which can donate a phosphate to ADP (in the next step) to form ATP. That this is an oxidation reaction should be obvious from the fact that the carbonyl C in G3P has two bonds to O but 3 bonds in 1,3 BPG. To carry out an oxidation reaction, you need an oxidizing agent. In comes NAD+, a modest but very prevalent oxidizing agent in biology. When glucose is oxidized completely by O2 to CO2 during combustion, much energy is released so we can surmise that oxidation reactions, if carried out by powerful oxidizing agents like O2, proceed with a large negative ΔGo. For every oxidation reaction, the oxidizing agent is reduced. All reactions are potentially reversible so the products formed are new potential oxidizing and reducing agents. As in acid/base reactions, which proceed from stronger acid to weaker conjugate acid, redox reactions proceed from a stronger to a weaker oxidizing agent. For reaction 6, we must use tables of redox potentials to calculate the actual ΔGo. It turns out to be close to 0, which is great since in the same reaction, a substrate-level phosphorylation reaction (using inorganic phosphate - Pi instead of ATP) occurs. In summary, this reaction catalyzes the first and only oxidation of glucose in glycolysis which has paid (thermodynamically) for the generation of a mixed anhydride whose phosphorylating potential is higher than that of ATP. Figure \(32$ shows a proposed mechanism for glyceraldehyde-3-phosphate dehydrogenase from Trypanosoma cruzi Figure $32$: Proposed mechanism for glyceraldehyde-3-phosphate dehydrogenase from Trypanosoma cruzi (after Reis et al. Phys. Chem. Chem. Phys., 2013, 15, 3772. https://pubmed.ncbi.nlm.nih.gov/23389436/) The reaction proceeds in two parts. The first (top section) is the oxidation of glyceraldehyde-3-phosphate (G3P) by NAD+ to the state of a thioester attached to Cys166. The now reduced NADH dissociates and is replaced by a new NAD+ for another cycle of catalysis. In the meantime, inorganic phosphate, Pi, binds and reacts with the thioester to form 1,3-bisphosphoglycerate (1,3-BPG). The mechanism is not entirely clear. There are two phosphate binding sites, Pi (red) and Ps (purple) which interact with phosphate groups on the substrates. The oxidation part of the reactions appears to take place at the Pi site. Pi is composed of the side chains of Thr197, Thr199, and the 2-hydroxyl group of the ribose in NAD+. The Pi site in the T. cruzi enzyme consist of Thr226 and Arg249, Gly227, and Ser 247. The mechanism appears to involve a flip of the orientation of substrates and intermediates after the dissociation of NADH from the enzyme. Figure $33$ shows an interactive iCn3D model of the Trypanosoma cruzi glyceraldehyde-3-phosphate dehydrogenase with bound NAD and a 1,3-bisphosphoglycerate analogue (1QXS) Figure $33$: Trypanosoma cruzi glyceraldehyde-3-phosphate dehydrogenase with bound NAD and a 1,3-bisphosphoglycerate analogue (1QXS) . (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...WnsNSgmA3AJyw6 Only two monomers of the homotetramer are shown for clarity. The Pi site is shown in magenta and the Ps site is in purple. The phosphonic acid analog is shown in spacefill and NAD in sticks. The active site His194 and Cys166 are also shown in sticks and labeled. Reaction 7: 1,3 BPG + ADP + H+ → 3PG + ATP ΔGo= -4.5 kcal/mol (-19 kJ/mol) This reaction, catalyzed by phosphoglycerate kinase, is shown in Figure $34$ Figure $34$: Summary reaction - phosphoglycerate kinase It's finally happened! An ATP has been made for each of the two 1,3-BPG molecules derived from glucose. We've made back the ATP used in steps 1 and 3. A mixed phosphoanhydride bond is broken in 1,3 BPG as a phosphoanhydride bond is made in ATP. As the mixed phosphoanhydride has higher energy than its hydrolysis product compared to the phosphoanhydride in ATP, the reaction proceeds with a negative ΔGo A mechanism for the reaction is shown in Figure $35$. Figure $35$: Reaction mechanism for phosphoglycerate kinase Figure $36$ shows an interactive iCn3D model of human phosphoglycerate kinase in complex with ADP, 3PG, and magnesium trifluoride (2WZB). Figure $36$: Human phosphoglycerate kinase in complex with ADP, 3PG, and magnesium trifluoride (2WZB). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...w5yvu2VuGhmQXA 3-phosphoglycerate is shown in spacefill with CPK colors. ADP plus the adjacent MgF3 is a mimetic for the transition state for ATP synthesis. Hence this structure shows the products/transition state in a closed active site, which as we have seen before prevents spurious hydrolysis of 1,3-BPG or ATP. Figure $37$ shows an interactive iCn3D model of the alignment of the open form of human phosphoglycerate kinase (2XE7) with bound 3PG and ADP with the closed form with bound 3PG, ADP, and MgF3 (2WZB). Figure $37$: Alignment of the open form of human phosphoglycerate kinase (2XE7) with bound 3PG and ADP with the closed form with bound 3PG, ADP, and MgF3 (2WZB). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...n5FfsJeYitiWr9 Use the "a" key to toggle back between the open form (magenta) and closed forms (cyan). In the closed state most closely representing the bond transition state of ATP, the two lobes of the enzyme clamp together. Reaction 8: 3PG ↔ 2PG ΔGo= +1.1 kcal/mol (4.6 kJ/mol) This reaction, catalyzed by phosphoglycerate mutase (PGM), is shown in Figure $38$ Figure $38$: Summary reaction - phosphoglycerate mutase This isomerization reaction proceeds with little thermodynamic barrier. Its function is to locate the phosphate on C2 which on the next reaction (dehydration) will form a molecule whose phosphoryl transfer potential is greater than ATP. It seems so simple but the enzymes that catalyze this reaction are diverse and quite complicated from a mechanistic perspective. There are two types of PGMs, bisphosphoglycerate and monophosphoglycerate mutases that carry out three different reactions involving shuffling of phosphates from one position to another in 3C sugars or cleavage of a phosphate from a sugar • 3-phosphoglycerate ↔ 2-phosphoglycerate (reaction 8 of glycolysis) catalyzed by bisphosphoglycerate and monophosphoglycerate (the glycolytic enzyme) mutases • 1,3-bisphosphoglycerate ↔ 2,3-bisphosphoglycerate by bisphosphoglycerate mutases • 2,3-phosphoglycerate ↔ 3-phosphoglycerate + Pi by bisphosphoglycerate mutases Within the monophosphoglcyerate mutases, and more specifically the phosphoglycerate mutase (PGM) of glycolysis, there are two types • one that depends on the cofactor 2,3-phosphoglycerate. These are called cofactor-dependent phosphoglycerate mutase (dPGM) and are found in mammals, yeast, and some bacteria. They do not require metal ions. • one that does not depend on the cofactor 2,3-phosphoglycerate. These are called cofactor-independent phosphoglycerate mutase (iPGM) and are found in plants and some bacteria. These can only interconvert 3PGA and 2PGA. One family of enzymes in the class requires Mn2+ while the other requires Mg2+ or Zn2+. These enzymes are often structurally similar to alkaline phosphatases The cofactor-independent and cofactor-dependent monophosphoglycerates (such as the phosphoglycerate mutase of glycolysis) are very different structurally and mechanistically so we will look at both types of mechanisms. Within each type, the enzyme sequences are very conserved. Mechanism of cofactor (2,3-BPG) dependent phosphoglycerate mutase (dPGM) The reaction is much less complicated than the cofactor-independent PGM. In E. Coli, the reaction involves the transfer of the phosphate on the C3-OH to the nucleophilic nitrogen on histidine 8 (His 10 in other enzymes) in the active site to form a covalent pHis8 intermediate. The phosphate on pHis could then be transferred to the O on carbon C2 of the substrate.  An active His 181in E. Coli may also act as a general acid/base and is adjacent to the Glu 88 in the active site. Figure $39$ shows an interactive iCn3D model of yeast phosphoglycerate mutase (cofactor dependent) bound to 3-phosphoglycerate (1QHF) Figure $39$: Yeast phosphoglycerate mutase (cofactor dependent) bound to 3-phosphoglycerate (1QHF). (Copyright; author via source). A dimer of the active tetramer is shown.  Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...9F4oc6mmr5DFs9 Mechanism of cofactor-independent phosphoglycerate mutase (iPGM) Let's consider the mechanism for the Mn2+-requiring iPGM from Geobacillus stearothermophilus. Two Mn2+ ions are in the active site. The mechanism for the first half of the cofactor-independent phosphoglycerate mutase (iPGM) reaction is shown in Figure $40$. Figure $40$: Part A - Mechanism for cofactor independent phosphoglycerate mutase (iPGM) from Geobacillus stearothermophilus (after Bolt et al, ibid) Arg 261 interacts with the substrate, stabilizing the negative charge in it and its transition state. It also makes the target phosphorous more electrophilic. Ser62 is activated by a Mn2+ ion to become more nucleophilic on the abstraction of a proton by Lys336. Reaction 3 probably proceeds through an SN2 mechanism. The rest of the mechanism is shown in Figure $41$. Figure $41$: Part B - Mechanism for cofactor independent phosphoglycerate mutase (iPGM) from Geobacillus stearothermophilus (after Bolt et al, ibid) The iPGMs are monomers with two distinct domains (lobes) containing a substrate binding site separated from an active site. They are connected by flexible sequences that bend to produce either an open or closed form of the enzymes (as we have seen before). In some enzymes, the two sites are merged at the interface between the domain. The active site Ser62 becomes phosphorylated and then transfers its phosphate to the new site in the substrate which is oriented differently in the enzyme. Figure $42$ shows an interactive iCn3D model of the Geobacillus stearothermophilus cofactor-Independent Phosphoglycerate Mutase (iPGM) with bound 2-phosphoglycerate (product) (1O98) Figure $42$: Cofactor-Independent Phosphoglycerate Mutase with bound 2-phosphoglycerate (product) (1O98). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...71iYHXAEKMGsEA Reaction 9: 2PG ↔ PEP + H2O ΔGo= +0.4 kcal/mol (+1.7 kJ/mol) This reaction, catalyzed by enolase, is shown in Figure $43$ Figure $43\: Summary reaction - enolase Now you can see the rationale for reaction 8. With a simple dehydration, a molecule with high phosphoryl transfer potential has been produced which in the next and final step of glycolysis produces ATP. This enzyme has an active site Mg2+ that is required for catalysis. Mammals have three forms of the enzyme, α-enolase (ENO-1) found in most tissues, β-enolase (ENO-3) found mostly in muscle, and γ-enolase (ENO-2) found in neurons. A possible mechanism for yeast enolase is shown in Figure \(44$. Figure $44$: A possible mechanism for yeast enolase Figure $45$ shows an interactive iCn3D model of Yeast enolase with bound 2-phosphoglycerate (7ENL) Figure $45$: Yeast enolase with bound 2-phosphoglycerate (7ENL). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...TgwYV9vF8cYtPA Reaction 10: PEP + ADP → Pyr + ATP ΔGo= -7.5 kcal/mol (-31 kJ/mol) This reaction, catalyzed by pyruvate kinase, is shown in Figure $46$ Figure $46$: Summary reaction - pyruvate kinase In this step, 1 more ATP is made for each PEP consumed (hence 2 ATPs for both 3C PEPs). The phosphoryl transfer potential for PEP is higher than for ATP, which allows this reaction to proceed with a large negative ΔGo (-7.5 kcal/mol, -31 kJ/mol). The mechanism for rabbit pyruvate kinase is shown in Figure $47$. Figure $47$: Mechanism for rabbit pyruvate kinase Figure $48$ shows an interactive iCn3D model of rabbit muscle pyruvate kinase complexed with Mn2+, K+, and pyruvate (1PKN) Figure $48$: Rabbit muscle pyruvate kinase complexed with Mn2+, K+, and pyruvate (1PKN). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...cZkm8DdEw2N3C6 We are done! Given that glycolysis is the central anaerobic energy-extracting pathway in all life, it is important that we examined each enzyme in detail. This is the net reaction of the glycolytic pathway: $\ce{Glc + Pi + 2ADP + 2NAD^{+} -> 2 Pyr + 2 ATP + 2NADH + 2H^{+} + 2H2O} \nonumber$ Uncoupling glycolytic oxidation and phosphorylation (ATP formation) Since we are most interested in energy transduction at this point, let's consider just two important steps in glycolysis that directly lead to ATP synthesis. Only one oxidative step is found in this pathway, namely the oxidative phosphorylation of the 3C glycolytic intermediate glyceraldehyde-3-phosphate, to 1,3-bisphosphoglycerate, a mixed anhydride (see link below for mechanism). The oxidizing agent is NAD+ and the phosphorylating agent is NOT ATP but rather Pi. The enzyme is named glyceraldehyde-3-phosphate dehydrogenase. It contains an active site Cys, which helps explain how the enzyme can be inactivated with a stoichiometric amount of iodoacetamide. A general base in the enzyme abstracts an H+ from Cys, which attacks the carbonyl C of the glyceraldehyde, forming a tetrahedral intermediate. Instead of the expected reaction (which would be the protonation of the alkoxide in an overall nucleophilic addition reaction at the aldehyde), a hydride leaves from the former carbonyl C to NAD+ in an oxidation step. Notice, this is a two-electron oxidation reaction similar to that seen in alcohol dehydrogenase. An acyl-thioester intermediate has formed, much like the acyl intermediate that formed in Ser proteases. Next inorganic phosphorous, Pi, attacks the carbonyl C of the intermediate in a nucleophilic substitution reaction to form the mixed anhydride product, 1,3-bisphosphoglycerate. Although we have formed a mixed anhydride, we cleaved a sulfur ester, which is destabilized with respect to its hydrolysis products (since the reactant, the thioester, is not stabilized by resonance to the extent of regular esters owing to the poor donation of electrons from the larger S to the carbonyl-like C.) In the next step, catalyzed by the enzyme phosphoglycerate kinase, ADP acts as a nucleophile that attacks the mixed anhydride of the 1,3-bisphosphoglycerate to form ATP. Note that the enzyme is named for the reverse reaction. We have coupled the oxidation of an organic molecule (glyceraldehyde-3-phosphate) to phosphorylation of ADP through the formation of a "high" energy mixed anhydride, 1,3-bisphosphoglycerate. The linkage between the oxidation of glyceraldehyde-3-phosphate and the phosphorylation of ADP by 1,3-bisphosphoglycerate can be artificially uncoupled by adding arsenate, which has a similar structure as phosphate. The arsenate can form a mixed anhydride at C1 of glyceraldehyde-3-phosphate, but since the bringing O-As bond is longer and not as strong as in the mixed anhydride, it is easily hydrolyzed. This prevents the subsequent transfer of phosphate to ADP to form ATP. Figure $49$ shows a summary of oxidation and substrate-level phosphorylation in glyceraldehyde-3-phosphate dehydrogenase. Figure $49$: Oxidation and substrate-level phosphorylation in glyceraldehyde-3-phosphate dehydrogenase (after Voet and Voet) Summary: Under anaerobic conditions, glucose (6Cs) is metabolized through glycolysis which converts it to two molecules of pyruvate (3Cs). Only one oxidation step has been performed when glyceraldehyde 3-phosphate is oxidized to 1,3-bisphosphoglycerate. To regenerate NAD+ so glycolysis can continue, pyruvate is reduced to lactate, catalyzed by lactate dehydrogenase. These reactions take place in the cytoplasm of cells actively engaged in the anaerobic oxidation of glucose (muscle cells for example during sprints). Note that t50}\). Figure $50$: Conversion of pyruvate to lactate We will explore the fate of pyruvate under anaerobic conditions more in the next chapter section.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/13%3A_Glycolysis_Gluconeogenesis_and_the_Pentose_Phosphate_Pathway/13.01%3A_Glycolysis.txt

Search Fundamentals of Biochemistry Introduction to Fermentation Fermentation is a process in which a fuel molecule is broken down anaerobically (or without oxygen) to meet ATP demands. One of the most notable pathways that can utilize fermentation is glycolysis, which we have just described. During glycolysis, glucose is converted to pyruvate, yielding a total of 2 ATP energy-rich molecules. This occurs in anoxic conditions (or without the requirement of oxygen). However, the process of glycolysis cannot be sustained if the end product is pyruvate. This is because the production of pyruvate also yields 2 molecules of reduced NADH in addition to the production of the two ATP molecules. The pool of available NAD+/NADH is limited within the body, and thus, this electron acceptor/donor molecule must continually be recycled for metabolic pathways to remain functional. For example, if NADH is not oxidized back into NAD+ in a timely manner, the glycolytic pathway can slow. Therefore, to be able to maintain the process of energy generation through glycolysis, the pool of NADH needs to be converted from the reduced form, back into the oxidized form (NAD+) form where it can then accept electrons from intermediates in the glycolytic pathway. This process of recycling NADH to NAD+ can occur aerobically through the Citric Acid Cycle, which is described in more detail in Chapter 16, or it can be processed via anaerobic fermentation (Figure \(1\)). Figure \(1\): Fates of Pyruvate via Aerobic or Anaerobic Pathways. In the 1860s, Louis Pasteur first described fermentation very narrowly, as the process that yeast use to convert glucose into ethanol when they are grown in the absence of air. Subsequently, it was discovered that other microorganisms could convert pyruvate to lactate (or lactic acid), instead of ethanol, during the process of anaerobic respiration. These microorganisms became known as lactic acid bacteria and are currently utilized heavily within the food industry to produce a wide array of fermented food products including cheese, yogurt, and sauerkraut, to name a few. As the enzymes within the glycolytic pathway were discovered, it became apparent that muscle tissue could also engage in anaerobic respiration-producing lactate. Thus while fermentative processes are most often described in microbial organisms, the definition of fermentation has since been broadened to include any enzymatic, energy-yielding pathways that occur in the absence of oxygen, including the production of lactate in muscle tissue of animals during the glycolytic process. Thus, NADH recycling to NAD+ typically occurs in anaerobic systems by two different routes: ethanolic fermentation or lactate fermentation (Figure \(1\)). We will focus on the details of these two systems here. Lactate Fermentation In lactate fermentation, pyruvate is converted to lactate by the enzyme lactate dehydrogenase. In the process, NAD+ is regenerated. The reaction catalyzed by lactate dehydrogenase (LDH) is shown in Figure \(2\). Figure \(2\): Reaction catalyzed by lactate dehydrogenase It is named for the reverse reaction which, as with other dehydrogenases, uses NAD+ as an oxidizing agent. The reaction is reversible with the ΔG0 for conversion of pyruvate to lactate of -3.76 kcal/mol (-15.7 kJ/mol). Under anaerobic conditions when glycolysis is the major source of ATP, pyruvate levels increase, further driving the reaction towards lactate formation and NAD+ regeneration so glycolysis can continue. The enzyme is found in the cytoplasm but a mitochondrial form also exists. It is most abundant in muscle, liver, kidney, and also in erythrocytes. Interestingly, mature red blood cells are enucleated and do not contain any mitochondria. Their lifespan is limited to approximately two weeks. During this time, their primary energy resources are generated through the process of anaerobic fermentation via the glycolysis-lactate pathway. The active enzyme is a tetramer of various compositions of two different subunits, the heart (H) and muscle (M) forms. The quaternary structures consist of 5 different isozyme forms containing the H and M subunits. LDH-1, found most abundantly in the heart, is a tetramer of 4H subunits (H4). The other forms are as follows: LDH-2 (H3M, prevalent in RBCs), LDL-3 (H2M2, prevalent in lungs), LDH-4 (HM3, prevalent in the kidney), and LDH-5 (M4, prevalent in muscle). During anaerobic metabolism, lactate is produced by muscle tissue and released into the bloodstream where it can travel back to the liver. Once in the liver, lactate is converted back into pyruvate and can be utilized to produce glucose through a pathway called gluconeogenesis. The liver can then export the glucose into the blood from where it can be taken up by the muscle for ATP production. This cycle is called the Cori cycle and is illustrated in Figure \(3\). Figure \(3\): Cori cycle Upper diagram is a cartoon image of the Cori cycle and the lower diagram demonstrates the recycling of NAD+/NADH in different locations within the body. Figure modified from Servier Medical Art Notably, anaerobic metabolism can only be sustained for short periods in animals due to its high energy demand. Aerobic respiration is required to maintain an adequate ATP supply. However, the production of lactate by certain tissues, such as white, or fast-twitch muscle cells occurs regularly and releases lactate into the bloodstream, where it can be taken up and used as an energy source by other neighboring tissues such as red, slow-twitch muscle. Brain tissue can also effectively use lactate as an energy source as well. Within these 'consumer' tissues, lactate is converted back into pyruvate using the mitochondrial LDH enzyme where it can then be processed by aerobic respiration producing high levels of ATP. Thus, the production of lactate can be thought of as a strain response that occurs during times of metabolic stress, such as intense cardiovascular exercise. It has also been noted that after an injury or head trauma, the activation of epinephrine will cause an increase in lactate production and blood levels of lactate will increase. Thus, it has been hypothesized that lactate may play a role in the repair of damaged tissue. Clinical trials and experiments are currently underway to determine if lactate can help in the healing and recovery process for conditions such as traumatic brain injury, myocardial infarction, and sepsis. Early studies have shown that lactate can increase the production of Brain-Derived Neurotropic Factor (BDNF) which supports neuronal growth, providing further support for the role of lactate in recovery and repair. Spectroscopy analyses show the geometry and electrostatics of the active sites of LDHs from all organisms are essentially the same. The mechanism for the reverse oxidation of lactate by NAD+ by spiny dogfish lactate dehydrogenase (LDH) is shown in Figure \(4\) Figure \(4\): Mechanism for the reduction of lactate by NAD+ by spiny dogfish lactate dehydrogenase (LDH) Lactate is shown with a deuterium (D), which moves as a deuteride to NAD+ to form NADH, simply to illustrate the stereochemistry of the reaction. In the reverse reaction, the reduction of pyruvate to lactate (to regenerate more NAD+ so anaerobic glycolysis can continue), the deuterium (or the proR H of non-deuterated NADH) is removed. Figure \(5\) shows an interactive iCn3D model of dogfish M4 apo-lactate dehydrogenase (1LDM). Figure \(5\): Dogfish M4 apo-lactate dehydrogenase (1LDM).(Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...FBzuqJ2gEPUHM8 Only the monomer of the active tetramer is shown. NAD and oxamic acid, structurally similar to lactate/pyruvate and a competitive inhibitor of the enzyme, are labeled. His193, the active site general base/acid, is near Arg 169 and to Asp 166 and Asn 138, to which it is hydrogen bonded. LDH exists in two major conformational states, T (inactive) and R (active) state, as we have seen with other allosteric proteins. Figure \(6\) shows an interactive iCn3D model of the T state (2ZQY) and R state (2ZQZ) of Lacticaseibacillus casei L-lactate dehydrogenase. Figure \(6\): T state (2ZQY) and R state (2ZQZ) of Lacticaseibacillus casei L-lactate dehydrogenase. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...zjrtVYjLXdhDp8 Toggle between the magenta T state and cyan R state using the "a" key. Orient the proteins to see the best view of the allosteric changes caused by binding regulators. The spheres show Arg 171 whose disposition changes significantly in the T and R states. Pyruvate (a reactant/product) activates the enzyme (conversion of T to R state) and is considered a homotropic regulator since it is a substrate. This is similar to the allosteric "activation" of hemoglobin by its ligand O2, which preferentially binds to the R state and promotes the T-to-R state transition. Fructose 1,6-bisphosphate, a glycolytic intermediate, also activates the Lacticaseibacillus casei L-lactate dehydrogenase and since it is not a reactant/product of the enzyme, it is considered a heterotropic allosteric regulator. The enzyme hence is effectively regulated by the concentration of substrate (pyruvate) and by NADH levels. High concentrations of ethanol consumption lead to high levels of NADH through the activity of alcohol dehydrogenase. High levels of NADH would lead to increased lactate production as well. Ethanol Fermentation in Yeast In this process, pyruvate is decarboxylated first to acetaldehyde by the thiamine pyrophosphate (TPP)-requiring enzyme pyruvate decarboxylase. The resulting product, acetaldehyde, is then reduced by NADH to form ethanol by the enzyme ethanol dehydrogenase in a process that reforms NAD+. Yeast are facultative (not obligate) anaerobes in that they can produce energy by glycolysis and ethanol fermentation in the absence of oxygen Figure \(7\):. Of course, in the presence of oxygen, the pyruvate produced from glycolysis in yeast is preferentially converted to acetyl-CoA which enters the citric acid cycle and oxidative phosphorylation pathways to maximize ATP production. Figure \(7\): Summary of Ethanol Fermentation in Yeast Figure from Thomas Baldwin Pyruvate decarboxylase Pyruvate decarboxylase catalyzes the first step in the alcoholic fermentation pathway converting pyruvate into acetaldehyde and carbon dioxide. Interestingly, in addition to yeast, some species of fish, such as goldfish and carp, have a homologous enzyme that allows for the production of ethanol when oxygen is scarce. Pyruvate decarboxylase is tetrameric occurring as a dimer of dimers with two active sites shared between the monomer subunits of each dimer. The decarboxylation reaction requires two cofactors, thiamine pyrophosphate (TPP) and magnesium (Figure \(8\)). Within the active site, the acidic Glu-477 and Glu-51 residues and the Mg2+ cofactor interact with and stabilize the TPP cofactor. The aminopyrimidine ring on TPP acts as a base and enables the formation of the TPP nucleophile with the removal of the C2 proton. This reaction is stabilized by the protonation of Glu-51. The nucleophilic attack of pyruvate causes the release of carbon dioxide. The acetaldehyde intermediate is still covalently attached to the TPP cofactor following the release of carbon dioxide. Figure \(8\): Part 1 Pyruvate Decarboxylase mechanism - decarboxylation Once carbon dioxide diffuses away from the active site, the double bond of the enol-intermediate abstracts a proton from Asp-28, which is stabilized by a neighboring His-115 residue. TPP can then serve as a good leaving group during the formation of the carbonyl functional group and causes the release of acetaldehyde (Figure \(9\):). Figure \(9\): Part 2 Pyruvate Decarboxylase mechanism - acetaldehyde generation Figure \(10\) shows an interactive iCn3D model of the thiamin diphosphate-dependent enzyme pyruvate decarboxylase from the yeast Saccharomyces cerevisiae (1PVD). Figure \(10\): Thiamin diphosphate-dependent enzyme pyruvate decarboxylase from the yeast Saccharomyces cerevisiae (1PVD).(Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...DMM4gjodX4q1G9 TTP is shown in spacefill. The enzyme is a homotetramer. Alcohol dehydrogenase (ADH) Alcohol dehydrogenase enzymes catalyze the interconversion of aldehydes or ketones with alcohol functional groups. In humans and other animals, they are utilized to break down alcohols that are ingested, as well as participate in the biosynthesis of numerous metabolites. There are at least 5 classes of alcohol dehydrogenases that use NAD+ as an oxidizing agent to convert alcohols to aldehydes or ketones. We are mainly concerned with the yeast ADH, a member of Family I, that is involved in ethanol fermentation. Figure \(11\) shows the reaction catalyzed by alcohol dehydrogenase. Figure \(11\): Reaction catalyzed by alcohol dehydrogenase Figure \(\PageIndex{12A}\) below shows a mechanism for yeast alcohol dehydrogenase (ADH1). ADH1 requires zinc as a metal cofactor and utilizes NADH as the reducing agent in the conversion of acetaldehyde into ethanol. Figure \(\PageIndex{12A}\): Simplified catalytic mechanism of ADH reactions. Upon binding of both substrates (e.g., NAD(P)+ and alcohol; 0->1->2) a hydride transfer occurs from the alcohol-carbon atom to the oxidized nicotinamide moiety yielding the Zn-coordinated carbonyl product and NAD(P)H (3). Both can dissociate from the active site yielding apo-ADH (0). Alternatively, only NAD(P)H stays bound and the reduced ADH can undergo a reductive conversion (1’->2’-> 3’-> 1). Amanda Silva de Miranda, et al. Front. Catal., 10 May 2022. Sec. Biocatalysis. https://doi.org/10.3389/fctls.2022.900554. Creative Commons Attribution License (CC BY). Figure \(\PageIndex{12B}\) shows key amino acids in the active site of yeast ADH1. Note that in this figure, two cysteines are the coordinating ligands. Figure \(\PageIndex{12B}\): Abbreviated mechanism for yeast alcohol dehydrogenase (ADH1) If you look at the reverse reduction reaction, the acetaldehyde (or a ketone for 20 alcohols) carbonyl is sp2 hybridized and planar, so the hydride could be added to either of the two faces, re or si, of the plane (using the same rules used to define R or S enantiomers). Likewise, the methylene carbon atom with two hydrogens, one of which is transferred as a hydride, is prochiral. Hence four different stereochemical hydride transfer pathways are possible, as shown in Figure \(\PageIndex{13A}\) below. Hydride transfers from the si-face of the prochiral aldehyde/ketone result in (R)-configured alcohols whereas hydride attacks from the re-face yield (S)-alcohols. In both cases, the hydride transferred can stem from either the re- or si-face of the nicotinamide ring. Figure \(\PageIndex{13A}\): Possible stereochemical courses of the hydride transfer from NAD(P)H to the ketone. Attacks from the si-face of the ketone result in (R)-alcohols (E1 and E2) whereas hydride attacks from the re-face (E3 and E4) result in (S)-alcohols. ADHs catalyzing hydride addition from the re-face of the ketone (or abstraction of a hydride from (S)-alcohols) are termed Prelog-selective ADHs whereas those ADHs attacking from the si-face (or abstracting a hydride from the (R)-alcohol) are termed anti-Prelog ADHs. Different ADHs are available for the different reaction stereochemistries. The stereochemistry for the reaction of ADH1 with ethanol is shown in Figure \(\PageIndex{13B}\) below. Deuterium (D) is shown in the figure to better illustrate the stereochemistry of the reaction. With the deuterium label, the carbon is now chiral and the enantiomer shown is the R isomer. The corresponding H in the undeuterated and prochiral form of ethanol is removed by the enzyme, as shown in Figure \(\PageIndex{13B}\) below. Figure \(\PageIndex{13B}\): Stereochemistry of human alcohol dehydrogenase reaction. Figure \(14\) shows an interactive iCn3D model of the Yeast alcohol dehydrogenase (ADH I) with bound substrate analogs- ADH1 (4W6Z). Figure \(14\): Yeast alcohol dehydrogenase structure with bound substrate analogs (4W6Z). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...feE7toF6wU6MB9 All four monomers of the homotetramer are shown. The light gray subunit has the key catalytic and binding residues shown in sticks and labeled. The numbering for the key residues is a bit different than in the mechanism shown above. The cofactor analog, nicotinamide-8-iodo-adenine-dinucleotide, is shown in sticks and the substrate analog, trifluoroethanol is shown in spacefill. Only two subunits (a homodimer) have both bound cofactor and substrate. The substrate trifluoroethanol in these subunits is ligated to the catalytic Zn2+ through its oxygen, with the other ligands provided by the side chains of Cys 43, Cys 153, and His 66. The other two monomers (the other homodimer) have a different conformation and use Glu 67 to coordinate the Zn2+ instead of His 66 and no substrate is bound. This may be an intermediate in the process that displaces water bound to Zn2+ with the substrate. Alcohol Metabolism in Humans: When ethanol is consumed, it is oxidatively metabolized primarily in the liver. Cytosolic Alcohol Dehydrogenase (ADH) and mitochondrial Acetaldehyde Dehydrogenase 2 (ALDH2) are the main enzymes involved in this metabolic pathway, first converting ethanol to acetaldehyde and then acetaldehyde to acetate (Figure \(15\):). Liver mitochondria have a limited capacity to use the acetate in the Kreb cycle because the enzyme needed to convert acetate to acetyl-CoA (acetyl-CoA synthase 2) is almost absent in the liver, but is abundant in the heart and skeletal muscles. Thus, most of the acetate resulting from ethanol metabolism escapes the liver into the blood circulation and is eventually metabolized to CO2 by way of the Kreb cycle in cells with mitochondria that contain enzymes to convert acetate to acetyl CoA, such as heart, skeletal muscle, and brain. Figure \(15\): Summary of Ethanol Metabolism in Humans Figure modified from Zakhari, S. (2013) Alcohol Research: Current Reviews 35(1):6-16 Like many substances, the consumption of alcohol can be both beneficial and detrimental, depending on the quantity and frequency of consumption. This is known as a hormetic response (Figure \(16\):). At low doses (up to 2 daily drinks for men and 1 daily drink for women) the consumption of alcohol can be cardioprotective. At higher consumption rates this protective effect is lost and the detrimental effects of alcohol consumption become apparent and include cardiotoxicity, liver damage, and increased cancer risk. Not to mention the debilitation that can accompany addiction. Figure \(16\): Alcohol Consumption and Hormesis The graph presented in Figure \(17\) exemplifies the hormetic nature of alcohol consumption. Maintaining ethanol consumption between one (for women) and two (for men) daily drinks will significantly reduce the overall risk of mortality. However, mortality risk skyrockets at high ethanol consumption. Figure \(17\): Risk of Mortality and Alcohol Consumption Moderate alcohol use of no more than 2 drinks per day for men, and no more than 1 drink per day for women can reduce the risk of coronary artery disease by 29%, and reduce the risk of dying from any cardiovascular disease by 25%. These statistics are significant. Do note that at these levels that there is an increased risk of dying from hemorrhagic stroke. Increased alcohol consumption, however, can have dire effects on health and can lead to unwanted addiction. The Centers for Disease Control estimates that alcohol abuse leads to approximately 80,000 deaths annually in the United States and that up to 40% of deaths related to liver disease are caused by alcohol abuse. Harvard Medical School has evaluated thousands of studies and has tried to come up with a method for alcohol risk assessment. They categorize low-risk drinking behavior for men as no more than four drinks in a single day or 14 drinks in a week and no more than 3 drinks in a single day or a total of 7 drinks in a single week for women. Women have lower drinking tolerance due to their smaller sizes, but also due to metabolic differences. Women have less ADH and typically higher fat levels which disperse and retain ethanol longer. Addiction and damage to the liver tend to occur more quickly in women as well. Harvard ranks people at increased risk if their drinking is above either the single day or the weekly limit (which is estimated at 29% of drinkers). High-risk drinkers break both the daily and weekly limits (estimated at 14% of drinkers). During ethanol metabolism, when circulating ethanol is in the millimolar range, acetaldehyde is in the micromolar range, and acetate is in the millimolar range (Figure \(18\):). When heavy drinking or chronic drinking occurs the Acetaldehyde Dehydrogenase enzyme cannot keep up with the Alcohol Dehydrogenase enzyme and the pool of Acetaldehyde increases. This aldehyde has many toxic effects within biological systems (Figure \(19\):). Figure \(18\): The Effects of Binge or Chronic Drinking on Ethanol Metabolites. Figure modified from Zakhari, S. (2013) Alcohol Research: Current Reviews 35(1):6-16 Acetaldehyde can form adducts with proteins. For example, acetaldehyde adducts on cytoskeletal components such as microtubulin, lead to the swelling of hepatocytes (liver cells). If secreted from the cell, these protein adducts can also be recognized as foreign by the immune system and cause an autoimmune response causing further inflammation and damage to the liver. Acetaldehyde also causes oxidative damage to lipids and DNA and can alter mitochondrial function. Overall, the liver is stressed and unhappy when too much ethanol is consumed. Figure \(19\): Acetaldehyde Toxicity Figure modified from Servier Medical Art Heavy chronic drinking can also lead to epigenetic modifications that alter protein expression patterns within the cell. Due to the oxidation of alcohol to acetate, the metabolism of alcohol also leads to an increase in the NADH: NAD+ ratio altering carbohydrate metabolism. Heavy ethanol consumption promotes the formation of reactive oxygen species and can also promote apoptosis. Overall, heavy drinking is extremely hard on liver function. It can lead progressively from the formation of a fatty liver to liver cirrhosis and increased risk of liver cancer. The risk for several other types of cancer is also heightened with heavy alcohol use. Chronic heavy alcohol use can also alter gene expression, especially in the liver. Figure \(20\) is a schematic representation of DNA methylation, which converts cytosine to 5′methyl-cytosine via the actions of DNA methyltransferase (DNMT). DNA methylation typically occurs at cytosines that are followed by a guanine (i.e., CpG motifs). Within the liver, chronic heavy drinking reduces pools of S-adenosylmethionine (SAM) while increasing homocysteine and S-adenosylhomocysteine (SAH). SAH further inhibits DNA methyltransferases (DNMTs) by negative feedback inhibition, ultimately resulting in global hypomethylation of DNA. This hypomethylation leads to the inappropriate expression of many genes, especially within the liver tissue. Figure \(20\): Effects of Chronic Alcohol Use on DNA Methylation Figure modified from Zakhari, S. (2013) Alcohol Research: Current Reviews 35(1):6-16 Two genes that are upregulated due to this hypomethylation express the Catalase enzyme and the p450 oxidoreductase enzyme, CYP2E1. Both of these enzymes are involved in oxidative pathways of alcohol metabolism that produce the toxic acetaldehyde intermediate (Figure \(21\)). Expression of both of these proteins become more prevalent in chronic alcohol consumption or when blood alcohol levels are high, as in cases of binge or heavy drinking. The activity of these enzymes can also lead to the formation of reactive oxygen species that contribute to global cellular damage (lipid peroxidation, DNA damage, protein damage, etc). Figure \(21\): Oxidative Pathways of Alcohol Metabolism Gene regulation in other areas of the body is also affected in response to chronic heavy alcohol consumption. This is due to the production of acetate during the metabolic pathway of alcohol that is released from the liver into the bloodstream. In other areas of the body, acetate is converted to acetyl-CoA by the enzyme Acetyl-coenzyme A (acetyl-CoA) synthetase (AceCS). AceCS is activated by Sirtuin 1, also known as NAD-dependent protein deacetylase (SIRT1). Acetyl-CoA is used by histone acetyltransferase (HAT) to acetylate the lysine residues in histone proteins. Histone acetylation causes these proteins to release the bound DNA, allowing regions to be opened up for transcription (Figure \(22\)). Thus, higher levels of acetate promote histone acetylation and increased gene expression. Figure \(22\): The Effects of Chronic Alcohol Consumption on Histone Acetylation. Note in the diagram shown, that SIRT1 also deacetylates histones, resulting in gene silencing. Thus, SIRT1 is a sensor that balances gene activation and silencing in the cell based on the cell’s energy status. Alcohol metabolism results in acetate formation, which is used in extrahepatic tissues to produce acetyl-CoA, upregulating histone acetylation within those tissues. NOTES: AceCS1 = Acetyl-CoA synthase 1; ADH = alcohol dehydrogenase; ALDH = Aldehyde dehydrogenase.  Figure from Zakhari, S. (2013) Alcohol Research: Current Reviews 35(1):6-16 In addition to these effects, both ADH and ALDH utilize the cofactor nicotinamide adenine dinucleotide (NAD+), which is reduced to NADH; as a consequence, during ethanol oxidation the ratio NADH/NAD+ is significantly increased, altering the cellular redox state and triggering several adverse effects, related to alcohol consumption. Glycolysis and the Kreb Cycle are downregulated due to low NAD+ levels. This results in lower pyruvate levels, and lower conversion of pyruvate to acetyl-CoA and also causes a decrease in gluconeogenesis (ie there is not enough pyruvate to drive glucose production). Thus, the pyruvate that does form favors anaerobic conversion to lactate and can result in lactic acidosis or lowering of the blood pH levels. Low rates of gluconeogenesis can also contribute to hypoglycemia which can be seen during binge drinking. Hyperuricemia or an increase in blood levels of uric acid can also occur due, in part, to increased production of ketone bodies and lactic acid. Both the ketone bodies and lactate can compete with uric acid for excretion into the urine within the kidney. The uric acid gets retained and heightens blood levels. Ketone bodies typically form during periods of starvation when carbohydrate stores have been depleted. When the liver can no longer efficiently maintain blood glucose levels, it will break down fatty acids into ketone bodies and secrete these into the bloodstream (Figure \(23\)). Ketone bodies, such as acetoacetate, acetone, and D-beta-hydroxybutyrate (which isn’t a real ketone, but is still referred to as a ketone body) are released into the bloodstream to compensate for the reduced glucose levels. Brain, heart, and skeletal muscle tissue can utilize ketone bodies as an energy source and this is a good short term solution to starvation. However, the formation of lactate and ketone bodies can severely reduce blood pH levels and induce a life-threatening state known as ketoacidosis. In addition to starvation, heavy alcohol consumption can induce ketogenesis inappropriately. Figure \(23\): Formation of Ketone Bodies. Figure from Sav vas As noted before, high levels of lactate and ketone bodies within the bloodstream can result in dehydration and reduced excretion of uric acid, leading to hyperuricemia. Over time, high uric acid levels can cause uric acid to precipitate, especially in joints where it causes painful gout flare-ups (Figure \(24\)). Figure \(24\): Hyperuricemia and Gout.   Image from www.scientificanimations.com According to the American Cancer Society, excessive alcohol use can also increase the risk for several different forms of cancer. • Cancers of the mouth, throat, voice box, and esophagus: Alcohol use raises the risk of these cancers. Drinking and smoking together raise the risk of these cancers even more than drinking or smoking alone. This might be because alcohol can help harmful chemicals in tobacco get inside the cells that line the mouth, throat, and esophagus. Alcohol may also limit how these cells can repair damage to their DNA caused by the chemicals in tobacco. • Liver cancer: Long-term alcohol use has been linked to an increased risk of liver cancer. Regular, heavy alcohol use can damage the liver, leading to inflammation and scarring. This might raise the risk of liver cancer. • Colon and rectal cancer: Alcohol use has been linked with a higher risk of cancers of the colon and rectum. The evidence for this is generally stronger in men than in women, but studies have found the link in both sexes. • Breast cancer: Even a few drinks a week are linked with an increased risk of breast cancer in women. This risk may be especially high in women who do not get enough folate (Vitamin B12) in their diet or through supplements. Alcohol can also raise estrogen levels in the body, which may explain some of the increased risks. Cutting back on alcohol may be an important way for many women to lower their risk of breast cancer. So how does alcohol consumption contribute to increased risks of cancer? Alcohol may help other harmful chemicals, such as those in tobacco smoke, enter the cells lining the upper digestive tract more easily. This might explain why the combination of smoking and drinking is much more likely to cause cancers in the mouth or throat than smoking or drinking alone. In other cases, alcohol may slow the body’s ability to break down and get rid of some harmful chemicals. Alcohol might affect the body’s ability to absorb some nutrients, such as folate. Folate is a vitamin needed as a cofactor for enzymes involved in amino acid biosynthesis. Absorption of nutrients can be even worse in heavy drinkers, who often have low levels of folate. These low levels may play a role in the risk of some cancers, such as breast and colorectal cancer. Alcohol can raise the levels of estrogen, a hormone important in the growth and development of breast tissue. This could affect a woman’s risk of breast cancer. Too much alcohol can also add extra calories to the diet, which can contribute to weight gain in some people. Being overweight or obese is known to increase the risks of many types of cancer. Overall, heavy ethanol consumption produces a wide spectrum of hepatic lesions (Figure \(25\)). Fatty liver (i.e., steatosis) is the earliest, most common response that develops in more than 90 percent of problem drinkers who consume 4 to 5 standard drinks per day. With continued drinking, alcoholic liver disease can proceed to liver inflammation (i.e., steatohepatitis), fibrosis, cirrhosis, and even liver cancer (i.e., hepatocellular carcinoma). Heavy drinking can also damage other organs, such as the pancreas and the brain, and can raise blood pressure. It also increases the risk of heart disease and stroke. In pregnant women, alcohol use, especially heavy drinking, may lead to birth defects or other problems with the fetus. Figure \(25\): The Effects of Heavy Chronic Drinking on the Liver In 1951, the FDA approved disulfiram for the treatment of alcoholism in the US (Figure \(26\)). Initially, the drug was prescribed in very high doses, often as high as 3,000 mg per day. The high doses led to reports of severe reactions to alcohol, some of which were fatal. It is an inhibitor of the ALDH-2, which will lead to an even higher increase in acetaldehyde concentration if alcohol is consumed. Thus, alcoholics that are taking this drug to quit drinking need to maintain sobriety. If they drink while taking this drug, they will become very ill due to the accumulation of acetaldehyde. Overall, the use of this drug supports abstinence, which is often hard to realistically achieve. Figure \(26\): Disulfiram At one time it was thought that before prescribing disulfiram to people, patients should experience mixing the drug with alcohol in a supervised setting. Researchers felt it was important for the individuals to have full knowledge of what would happen if they mixed disulfiram and alcohol. This practice is no longer used, but every person must be educated on the reactions of combining alcohol with disulfiram before a prescription is written. Along with these effects, alcohol may contribute to cancer growth in other, unknown ways. and other mammals, various ADHs are used to oxidatively metabolize ethanol to acetaldehyde (also toxic) which is converted to acetate by the enzyme aldehyde dehydrogenase which catalyzes the following reaction: CH3CHO + NAD+ + H2O → acetate + NADH + H+ They can also convert methanol to the very toxic and reactive formaldehyde, which makes methanol poisoning so dangerous if not fatal. Mammalian enzymes are dimers of up to nine different monomers.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/13%3A_Glycolysis_Gluconeogenesis_and_the_Pentose_Phosphate_Pathway/13.02%3A_Fates_of_Pyruvate_under_Anaerobic_Cond.txt

Search Fundamentals of Biochemistry Introduction Gluconeogenesis is a metabolic pathway that results in the generation of glucose from non-carbohydrate carbon substrates such as lactate, glycerol, and glucogenic amino acids. It is one of the two main mechanisms humans and many other animals use to keep blood glucose levels from dropping too low (hypoglycemia). The other means of maintaining blood glucose levels is through the degradation of glycogen (glycogenolysis). Gluconeogenesis is a ubiquitous process, present in plants, animals, fungi, bacteria, and other microorganisms. In animals, gluconeogenesis takes place mainly in the liver and, to a lesser extent, in the cortex of the kidneys. This process occurs during periods of fasting, starvation, low-carbohydrate diets, or intense exercise and is highly endergonic. For example, the pathway leading from phosphoenolpyruvate to glucose-6-phosphate requires 6 molecules of ATP. Gluconeogenesis is often associated with ketosis. Gluconeogenesis is also a target of therapy for type II diabetes, such as metformin, which inhibits glucose formation and stimulates glucose uptake by cells. Lactate is transported back to the liver where it is converted into pyruvate by the Cori cycle using the enzyme lactate dehydrogenase. Pyruvate, the first designated substrate of the gluconeogenic pathway, can then be used to generate glucose (Figure \(1\)). All citric acid cycle intermediates, through conversion to oxaloacetate, amino acids other than lysine or leucine, and glycerol can also function as substrates for gluconeogenesis. Transamination or deamination of amino acids facilitates the entry of their carbon skeleton into the cycle directly (as pyruvate or oxaloacetate), or indirectly via the citric acid cycle. Glycerol, which is a part of the triacylglycerol molecule, can be used in gluconeogenesis. Gluconeogenesis is a pathway consisting of eleven enzyme-catalyzed reactions. The pathway can begin in the mitochondria or cytoplasm, depending on the substrate being used. Many of the reactions are the reversible steps found in glycolysis (Figure \(1\)). In humans, gluconeogenesis is restricted to the liver and a lesser extent the kidney. Figure\(1\): Gluconeognesis Bypass I (reverse of step 10 in glycolysis): Pyruvate to Phosphoenolpyruvate The conversion of pyruvate into phosphoenolpyruvate requires two enzymatic steps and the formation of oxaloacetate as the intermediate. In all species, the formation of oxaloacetate from pyruvate and any other TCA cycle intermediates is restricted to the mitochondrion, and the enzymes that convert PEP to glucose are found in the cytosol. The location of the enzyme that links these two parts of gluconeogenesis by converting oxaloacetate to PEP, PEP carboxykinase, is variable by species: it can be found entirely within the mitochondria, entirely within the cytosol, or dispersed evenly between the two, as it is in humans. Transport of PEP across the mitochondrial membrane is accomplished by dedicated transport proteins; however, no such proteins exist for oxaloacetate. Therefore in species that lack intra-mitochondrial PEP, oxaloacetate must be converted into malate or aspartate, exported from the mitochondrion, and converted back into oxaloacetate to allow gluconeogenesis to continue. Here is the net reaction: Pyruvate + ATP + HCO3- + GTP → PEP + ADP + GDP + CO2 ΔGo' = +0.2 kcal/mol (0.8 kJ/mol) Figure\(2\) shows the overall reaction for the conversion of pyruvate to phosphoenol pyruvate. Figure \(2\): Overall reaction for the conversion of pyruvate to phosphoenol pyruvate In systems that produce oxaloacetate in the mitochondria and then need to transport it to the cytosol to be converted into phosphoenolpyruvate, three enzymes are needed for the process: pyruvate carboxylase (PC) located in the mitochondrial matrix, aspartate aminotransferase (AAT), located in the matrix and the cytosol, and the phosphoenolpyruvate carboxykinase (PCK) located in the cytosol. Oxaloacetate and aspartate are intermediate compounds formed in the process. In the first reaction, pyruvate carboxylase converts pyruvate into oxaloacetate, based on the following reaction: Pyruvate carboxylase: pyruvate + HCO3- + ATP → oxaloacetate + ADP + Pi The pyruvate carboxylase enzyme requires biotin as a cofactor and has two major enzymatic functions: 1) carbon fixation from carbon dioxide, and 2) carbon transferase activity, placing the carbon dioxide that has been fixed, onto the molecule of pyruvate to create oxaloacetate. The biotin cofactor is shown in Figure (3\). Biotin is a water-soluble vitamin (known as D-biotin or vitamin B7) and is a cofactor in many enzymatic reactions, especially those involving carboxylation (or carbon fixation reactions). It is part of the vitamin B2 complex and is an essential vitamin for mammals. Deficiency results in dermatitis, loss of hair, and neurologic symptoms. Figure (3\): Chemical Structure of Biotin. Biotin is shown in several different formats, (A) line structure, (B) stick model, (C) ball and stick model, and (D) spacefilling model. Figure from: Biosynthesis The biotin cofactor forms an amide linkage with the pyruvate carboxylase enzyme at a lysine residue (Figure (4\)). This creates a flexible linker region within the pyruvate carboxylase that is capable of dipping the biotin cofactor into the different catalytic domains of the enzyme to obtain the biological activity of the protein. Attaching the biotin to the enzyme requires the energy of ATP. Figure (4\) Attachment of Biotin Cofactor to Lysine Residue in Pyruvate Carboxylase. Image from: Biosynthesis The pyruvate carboxylase enzyme is a tetramer that contains four functional protein subunits as shown in Figure \(5\). The BCCP flexible arm is capable of extending into the biotin carboxylation (BC) domain where biotin is first carboxylated, and then the BCCP shifts over to the carboxyl transferase (CT) domain where the fixed molecule of carbon dioxide can be transferred to pyruvate forming oxaloacetate. Figure \(5\) Structure of the Pyruvate Carboxylase Enzyme. The overall structure of the Pyruvate Carboxylase is a tetramer that contains four functional protein subunits. Two subunits are shown in color, with the other two indicated in gray at the back of the structure. (a) shows the space-filling model while (b) shows the major domains as a cartoon graphic. Each subunit contains a biotin carboxylase (BC) domain shown in blue, a carboxytransferase (CT) domain shown in yellow, and the biotin-carboxyl carrier protein (BCCP) domain shown in red and green. Figure modified from Liu, Y., et al (2018) Nat Commun 9:1384 In the first part of the reaction, the biotin cofactor is carboxylated using bicarbonate as a substrate with ATP to drive the formation of a molecule with high energy with respect to its hydrolysis product intermediate.  (Remember, there is no such thing as a "high energy" bond.) Thus, biotin is acting as an intermediate carrier of the carboxy group that will be added to pyruvate during the formation of oxaloacetate. This occurs in the biotin carboxylase domain (BC) with biotin bound to a biotin carboxyl carrier protein (BCCP) domain. The carboxyl group is then transferred to pyruvate to form oxaloacetate in the carboxyl transferase (CT) domain. Let's break up the mechanism of the enzyme from Rhizobium etli into three figures. Figure\(6\) below shows the first steps in the carboxylation of biotin in the biotin carboxylase domain, using bicarbonate as a substrate. In Figure\(6\), you can see that an oxygen from bicarbonate mediates nucleophilic attack on the outer phosphate group of the ATP molecule forming the carbonyl-phosphate intermediate + ADP. The carbonyl-phosphate decomposes to release carbon dioxide and phosphate. The negatively charged phosphate activates the biotin cofactor to enable a nucleophilic attack on the carbon of the carbon dioxide producing the carboxybiotin intermediate, shown in the upper part of Figure\(7\). Figure \(6\): The Carboxylation of Biotin during the Pyruvate Carboxylase Reaction Mechanism. Bicarbonate, shown in blue, enters the active site of the enzyme and reacts with the terminal phosphate of ATP forming a phosphorylated intermediate. This reaction is stabilized by two Mg2+ cofactors. The carboxyphosphate intermediate decomposes to release carbon dioxide and phosphate. Image modified from Ribeiro AJM et al. (2017), Nucleic Acids Res, 46, D618-D623. Mechanism and Catalytic Site Atlas (M-CSA): a database of enzyme reaction mechanisms and active sites. DOI:10.1093/nar/gkx1012. PMID:29106569. https://www.ebi.ac.uk/thornton-srv/m-csa/entry/223/. Creative Commons Attribution 4.0 International (CC BY 4.0) License. Figure\(7\): Carboxylation of biotin in pyruvate carboxylase. The upper diagram shows the actual carboxylation of biotin, which occurs in the BC domain. The carboxybiotin intermediate then shifts on the flexible arm and enters the carboxytransferase (CT) domain. Pyruvate enters the active site and this begins the steps involved in the carboxylation of pyruvate to form oxaloacetate. Ribeiro, ibid The carboxyltransferase activity is housed in the CT domain of the enzyme and requires the formation of a carbanion intermediate to occur on the pyruvate molecule to make it reactive enough to cause the addition of the carboxyl group. A base from the enzyme eliminates a proton from the C3 position of pyruvate causing the formation of an enol reaction intermediate. As the electrons on the oxygen from the pyruvate molecule reform the carbonyl group, the electrons from the carbon-carbon double bond shift onto the carbon, creating a reactive carbanion intermediate. At this point, the carboxybiotinyl complex swings into the CT domain, providing the carboxyl group that will be incorporated into pyruvate. The carbanion intermediate mediates a nucleophilic attack onto the carboxyl carbon from the BCCP complex, and carbon dioxide is transferred to the pyruvate, creating oxaloacetate. The biotin cofactor is restored by the donation of the proton from the basic residue in the enzyme. This also restores the Pyruvate Carboxylase enzyme for another round of activity. Figure\(8\) below shows the carboxylation of the enolate form of pyruvate to form oxaloacetate. Figure\(8\): Carboxylation of the enolate form of pyruvate to form oxaloacetate by pyruvate carboxylase. Ribeiro et al, ibid Figure\(9\) below shows an interactive iCn3D model of the biotin-dependent multifunctional enzyme pyruvate carboxylase from Rhizobium etli (2QF7) Figure\(9\): Biotin-dependent multifunctional enzyme pyruvate carboxylase from Rhizobium etli (2QF7). The enzyme is a homotetramer with C2 symmetry but only one subunit chain is shown for clarity. The biotin carboxylase domain (BC) is shown in cyan, the carboxytransferase domain (CT) in yellow, a C-terminal biotin carboxyl carrier protein (BCCP) domain (disconnected in parts) in magenta, and the allosteric domain in green. AGS in the cyan domain is a phosphothiophosphoric acid-adenylate ester, an ATP analog. It is shown interacting with the 2 Mg2+ ion and it is represented in sticks and labeled. Acetyl-CoA and a nonhydrolyzable analog, ethyl-CoA, are allosteric activators. CoASH (ethyl group not shown) is illustrated in spacefill bound to the green allosteric domain. Acetyl-CoA binding leads to a conformational change that decreases the distance between the two active sites on the BT (cyan) and CT (yellow) domains. Biotin, not shown, is tethered, in the BCCP domain, which transfers biotin between the two catalytic domains not within a single monomer (as shown above), but between different monomers in the homo 4-mer. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...7g91JmzJMad4r9 Figure\(10\) shows the flexible nature of the biotin flexible arm of the enzyme during the catalytic process. Figure\(10\) The Catalytic Activity of Pyruvate Carboxylase. The Biotin-Carboxyl Carrier Protein Domain (BCCP) is a flexible arm of the protein structure that can swing from one catalytic domain to the other during the reaction process. As shown in the diagram, the BCCP domain of subunit 1 will be fixed with carbon dioxide at the biotin carboxylase domain from subunit 1 (BC1). The BCCP will then translocate to the carboxyltransferase domain from subunit 2 (CT2) where it will create one molecule of oxaloacetate and reset the biotin cofactor. The BCCP will then swing into the BC2 domain where carbon fixation will occur. BCCP then shifts to the CT1 domain where it creates a second oxaloacetate molecule and resets the biotin. While this is going on with the BCCP from subunit 1, the other BCCP arms from subunits 2, 3, and 4 are also active. Thus, one full sequence of the enzyme activity will produce 8 molecules of oxaloacetate. Image from: Liu, Y., et al (2018) Nat Commun 9:1384 The next reaction in the process requires the conversion of oxaloacetate to phosphoenolpyruvate by phosphoenolpyruvate carboxykinase (PEPCK). However, the major human PEPCK enzyme, PCK1, resides in the cytoplasm of the cell. Thus the oxaloacetate produced in the matrix of the mitochondria by pyruvate carboxylase must be transported into the cytosol. However, there are no oxaloacetate transporters that can mediate the transfer directly. To leave the mitochondrial matrix, oxaloacetate has to first be converted to aspartate by the aspartate aminotransferase enzyme (AAT). It can then be transported to the intermembrane space of the mitochondria through an antiporter that transports one molecule of aspartate out of the matrix and one molecule of glutamate into the matrix Figure\(11\). Once in the intermembrane space, the aspartate can pass freely through a pore in the outer mitochondrial membrane. When it reaches the cytoplasm, aspartate is reconverted to oxaloacetate using the cytoplasmic aspartate aminotransferase enzyme (AAT). Oxaloacetate can then be used as a substrate by the phosphoenolpyruvate carboxykinase enzyme (PEPCK). Figure\(11\) Conversion of Oxaloacetate to Aspartate and Transfer to the Cytoplasm. Image modified from SMART Servier Medical Art The glutamate/aspartate transporter has some additional complexities associated with it. It cannot function on its own. It requires the coordinated functioning of the malate/alpha-ketoglutarate antiporter. Together, these antiporters are known as the Malate-Aspartate Shuttle System. The Malate-Aspartate Shuttle System is dependent on the functioning of two enzymatic processes. The first is the aspartate aminotransferase that was indicated more simplistically in Figure\(11\). The aspartate aminotransferase enzyme can utilize glutamate as an amine donor to generate aspartate from oxaloacetate. Alpha-ketoglutarate is also formed in this process. Depending on substrate concentrations and other regulatory mechanisms, this enzyme can also work in the reverse reaction to produce glutamate and oxaloacetate. In a different reaction using Malate Dehydrogenase, oxaloacetate can be reduced to form malate using a molecule of NADH as the electron donor. Figure\(12\) Chemical Reactions of the Malate-Aspartate Shuttle System Figure modified from Son, H.F. and Kim, J-J. (2016) PLoS One 10.1371 The malate dehydrogenase enzyme is expressed to high levels in the cytoplasm of liver cells as well as in the matrix of the mitochondria. This enzyme is a component of the Kreb Cycle where it mediates the formation of oxaloacetate in the last step of the cycle. Within the cytoplasm, it predominantly converts oxaloacetate to malate. The malate can then be shuttled into the matrix of the mitochondria through the malate/alpha-ketoglutarate antiporter (Figure\(13\)). In this antiporter, malate moves into the mitochondrial matrix while alpha-ketoglutarate moves into the intermembrane space (and subsequently into the cytosol of the cell). Together with aspartate, it can be used by cytoplasmic aspartate aminotransferase to produce glutamate and oxaloacetate. This is the oxaloacetate pool that is then utilized in the gluconeogenic pathway when PEPCK is active. The glutamate generated from this reaction is transported back into the matrix of the mitochondria through the aspartate/glutamate antiporter. The pool of aspartate in the matrix of the mitochondria is supplied by the reaction of the aspartate aminotransferase enzyme, which completes the reverse reaction from the one seen in the cytoplasm. In the matrix, aspartate aminotransferase uses glutamate and oxaloacetate as substrates to generate aspartate and alpha-ketoglutarate. This enables the transport of aspartate and glutamate through their specific antiporter. Within the gluconeogenic pathway, heightened levels of oxaloacetate are produced in the matrix of the mitochondria. Oxaloacetate is then converted to aspartate and transported across the inner membrane, where it can subsequently be converted back into oxaloacetate and used for the production of glucose. Figure\(13\): The Malate-Aspartate Shuttle System. Once oxaloacetate has been effectively transported via the Malate-Aspartate Shuttle into the cytoplasm, it is converted to phosphoenolpyruvate (PEP) by the PEP carboxykinase (PEPCK). The overall reaction mediated by PEPCK is: PEP carboxykinase (PEPCK): oxaloacetate + GTP → phosphoenol pyruvate + GDP PEPCK is classified as a lyase enzyme and exists in two isozymes, a cytoplasmic PEPCK, and a mitochondrial PEPCK. The cytoplasmic form is the one that is predominantly used in the gluconeogenic pathway and requires the action of the Malate-Aspartate Shuttle. However, small amounts of PEP can be made directly by the mitochondrial PEPCK and then transported across the mitochondrial membrane. The conversion of oxaloacetate to phosphoenolpyruvate by PEPCK mediates the removal of carbon dioxide and the addition of a phosphate group. A molecule of GTP is used during this process as the phosphate donor and magnesium ions serve as a cofactor. Figure \(14\) shows the reaction mechanism for the human PEP carboxykinase. Figure \(14\): Reaction mechanism for human PEP carboxykinase (PEPCK) Figure\(15\) below shows an interactive iCn3D model of phosphoenolpyruvate carboxykinase (PEPK) with a bound PEP and GTP-competitive inhibitor (1NHX). Figure \(15\): Phosphoenolpyruvate carboxykinase (PEPK) with a bound PEP and GTP-competitive inhibitor (1NHX). PEP is shown in colored sticks. The competitive inhibitor is shown in colored spacefill. The active site residues from the human PEPCK mechanism are shown as labeled sticks. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...QFcTssCJbuW2K7 Phosphatase Reactions of Gluconeogenesis (reverse of steps 1 and 3 in glycolysis): The last two unique enzymes of the gluconeogenic pathway are both phosphatase enzymes. Fructose 1,6-bisphosphatase converts fructose 1,6-bisphosphate to fructose 6-phosphate. The same phosphoglucoisomerase used in glycolysis can convert fructose 6-phosphate back into glucose 6-phosphate. A unique glucose 6-phosphatase enzyme will then convert glucose 6-phosphate to free glucose. The unique phosphatase reactions are shown in Figure\(16\). Figure \(16\): Phosphorylase Enzymes utilized during Gluconeogenesis First, let's consider the dephosphorylation of fructose 1,6-bisphosphate. The gluconeogenic enzyme is named fructose-1,6-bisphosphatase, (FBP or FBPase-1). FBP requires a metal cofactor and is competitively and allosterically regulated. Fructose 2,6-bisphosphate and low energy load (AMP and ADP) inhibit the enzyme. The overall reaction is shown below: Fructose-1,6-bisphosphatase: F1,6BP + H2O → F6P + Pi Figure \(17\) shows the mechanism of mouse fructose,1-6-bisphosphatase (FBPase-1) Figure \(17\): Mechanism of mouse fructose,1-6-bisphosphatase. Mg2+ ions coordinate the positioning of F-1,6-BP near an activated water molecule that mediates nucleophilic attack on the 1-position phosphate group. This leads to the cleavage of the phosphate group from the sugar molecule, F6P, which can then leave the active site. Figure\(18\) below shows an interactive iCn3D model of pig fructose-1,6-bisphosphatase with bound Mg2+, fructose-6-phosphate, and phosphate in the R state (1EYI). Figure \(18\): Fructose-1,6-bisphosphatase with bound Mg, fructose-6-phosphate and phosphate in the R state (1EYI). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...DbWMHBRTdPGhJ8 FBP is an allosteric homotetramer and can exist in a tense (T) and relaxed (R) state. When the enzyme is in the R state, it has the highest catalytic activity. Figure\(18\) shows only one monomer of the R state with bound ligands and substrates for clarity. Figure \(19\) shows an interactive iCn3D model of tetrameric FBP enzyme in the T state with bound Mg2+, fructose-6-phosphate, phosphate, and AMP, an allosteric inhibitor, in the T state (1EYJ). Figure \(19\): Tetrameric fructose-1,6-bisphosphatase in the T state with bound Mg2+, fructose-6-phosphate, phosphate and AMP in the T state (1EYJ). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...UAbjNtkPsoBdA9 FBPase is inhibited by AMP and fructose-2,6-bisphosphate, and activated by ATP.  Allosteric changes control the catalytic activity of the enzyme and the transition from the T to the R state, or vice versa. As shown in Figure\(17\), FBP has 3 metal binding sites. The mechanism shown in Figure \(17\) In the model, all of the binding sites are filled with Mg2+, however, Zn2+ can also play a role. In the presence of AMP, only one site is occupied by Mg2+, as shown in the model of the T state in Figure\(19\). The T state form of FBP has a loop (52-72) that is disordered. In the R state (no AMP), the loop interacts with the active site and 1 Zn2+ and 2 Mg2+ ions are bound in the three sites. The binding of AMP leads to the dissociation of two of the bound metal cofactors and causes a decrease in enzyme activity. The resulting fructose 6-phosphate is isomerized to glucose 6-phosphate. Glucose 6-phosphate is then converted by the last unique gluconeogenic enzyme, glucose 6-phosphatase, to generate free glucose. The overall reaction is shown below: Glucose-6-phosphatase: G6P + H2O → glucose + Pi The dephosphorylation of glucose only occurs appreciably in liver cells, as this is the primary location for the regulation of blood glucose levels. This serves as the final step in the gluconeogenic pathway. The glucose 6-phosphatase enzyme is a transmembrane protein that resides in the inner membrane of the endoplasmic reticulum. Thus, for glucose 6-phosphate to be dephosphorylated, it must first be transported from the cytoplasm into the lumen of the endoplasmic reticulum (ER) through transporter 1 (T1) (Figure\(20\)). The glucose 6-phosphatase (G-6-Pase) then cleaves the phosphate from the substrate, releasing inorganic phosphate (P) and glucose (red molecule). The inorganic phosphate is then transported back into the cytoplasm through transporter 2 (T2) and glucose is transported through Transporter 3 (T3). Free glucose is then transported back into the bloodstream through a glucose (GLUT) transporter (not shown in Figure\(20\)). Figure\(20\) Dephosphorylation of Glucose 6-Phosphate in the Lumen of the Endoplasmic Reticulum. The catalytic site of this hydrolase enzyme is comprised of Lys76, Arg83, His119, Arg170, and His176. His 176 acts as a nucleophile that attacks the phosphorous of G6P in an SN2-like reaction to form a His-PO32- intermediate. Hydrolysis follows restoring the enzyme to its original state. Figure \(21\) shows an abbreviated mechanism for the reaction. Figure \(21\): Abbreviate mechanism for human glucose-6-phosphatase Figure \(21\) shows an interactive iCn3D model of the AlphaFold predicted structure of human glucose-6-phosphatase (P35575). The enzyme contains nine transmembrane helices that dock the protein in the inner membrane of the ER. The active site faces the ER lumen. Figure \(14\): AlphaFold predicted structure of human glucose-6-phosphatase (P35575)(Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...jmcQAo49eZ8p16 Summary: It is noteworthy that the process of gluconeogenesis is energy intensive. For example, the phosphatase hydrolysis reactions that liberate glucose are not reverse kinase reactions and they do not produce substrate-level phosphorylation the way reactions occur in the second half of the glycolytic pathway. Overall, there are approximately 6 molar equivalents of ATP required to make 1 mole of glucose. Over 3 times as much energy is consumed during glucose formation than is generated from glycolysis during the breakdown of glucose. Per glucose molecule made, 2 ATP are used in the pyruvate carboxylase step, 2 GTP are used in the phosphoenolpyruvate carboxykinase step, and 2 ATP are used in the kinase reaction to create 1,3-bisphosphoglycerate from 3-phosphoglycerate. There is also the loss of 2 moles of NADH during the reverse reaction to create glyceraldehyde 3-phosphate from 1,3-bisphosphoglycerate, which reduces energy potential through oxidative phosphorylation in the mitochondria. Plus there are energy costs to changing the metabolite pools in the matrix of the mitochondria and for transporting molecules across the mitochondrial membrane that we have not considered here.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/13%3A_Glycolysis_Gluconeogenesis_and_the_Pentose_Phosphate_Pathway/13.03%3A_Gluconeogenesis.txt

Search Fundamentals of Biochemistry Introduction The pentose phosphate pathway (PPP), also known as the pentose phosphate shunt, is an important part of glucose metabolism. The PPP branches after the first step of glycolysis and consumes the intermediate glucose 6-phosphate (G6P) to generate fructose 6-phosphate (F6P) and glyceraldehyde 3-phosphate (G3P) through the oxidative and non-oxidative branches of the PPP. Unlike glycolysis and glucose aerobic oxidation, the PPP does not provide adenosine 5′-triphosphate (ATP) to meet the energy demands of cells. Instead, it supplies NADPH and ribose 5-phosphate (R5P). These two metabolites are vital for the survival and proliferation of cells. R5P is a building block for nucleic acid synthesis. NADPH is the reducing power required for the synthesis of fatty acids, sterols, nucleotides, and non-essential amino acids. Moreover, NADPH-derived conversion of oxidized glutathione (GSSG) to reduced glutathione (GSH) via glutathione reductase is important for cellular antioxidant defenses. Interestingly, NADPH also serves as the substrate of NADPH oxidases (NOXs) which produce reactive oxygen species (ROS). Both the oxidative branch and non-oxidative branch of the PPP take place in the cytosol (Figure \(1\)). Glucose 6-phosphate dehydrogenase (G6PD) is the rate-limiting enzyme of the oxidative PPP, determining the flux of G6P directed into the pathway. G6PD catalyzes the conversion of G6P to 6-phosphogluconolactone, accompanied by NADPH production. 6-phosphogluconolactonase (6PGL) is the enzyme that hydrolyzes 6-phosphogluconolactone to produce 6-phosphogluconate (6PG). 6-phosphogluconate dehydrogenase (6PGD) converts 6-PG to ribulose 5-phosphate (Ru5P) and generates NAPDH (Figure \(1\)). The largest contributor to cytosolic NADPH is the oxidative PPP in mammalian cells. The non-oxidative branch is composed of a series of reversible transfer reactions of chemical groups. Ribose 5-phosphate isomerase (RPI) and ribulose 5-phosphate epimerase (RPE) catalyze reversible reactions converting Ru5P to R5P and xylulose 5-phosphate (Xu5P), respectively. TKT catalyzes two reversible reactions. One is the conversion of Xu5P and R5P to G3P and sedoheptulose 7-phosphate (S7P). The other is the conversion of Xu5P and erythrose 4-phosphate (E4P) to G3P and F6P. Therefore, TKT can bi-directionally regulate the carbon flux between the non-oxidative PPP and glycolysis or gluconeogenesis. Transaldolase (TALDO) reversibly converts G3P and S7P to E4P and F6P. The non-oxidative branch not only replenishes metabolites of the oxidative branch (by their reversal), but also regulates the flux of glycolysis or gluconeogenesis by providing F6P and G3P (Figure \(1\)). Figure \(1\): The pentose phosphate pathway (PPP). The PPP branches after the first step of glycolysis and goes back to fructose 6-phosphate and glyceraldehyde 3-phosphate in the glycolytic and gluconeogenic pathway. The PPP produces R5P and NADPH for biosynthesis and redox regulation. Enzymes in the oxidative and non-oxidative PPP are shaded in green.  Figure from: Ge, T., et al. (2020) Cellular Endocrinology DOI:10.3389 Oxidative branch The oxidative branch of PPP (ox-PPP) is a non-reversible metabolic pathway where glucose-6-phosphate (G6P) is transformed into 6-phosphoglucono-δ-lactone by glucose-6-phosphate dehydrogenase (G6PD) and, subsequently, to ribulose-5-phosphate by 6-phosphogluconate dehydrogenase (6PGD) with the concomitant production of nicotinamide adenine dinucleotide phosphate (NADPH). The resulting ribulose-5-phosphate is then converted to ribose-5-phosphate and used for the biosynthesis of nucleotides (Figure\(2\)). Figure \(2\): Summary of the oxidative branch of phosphopentose pathway The first enzyme in the oxidative branch is the glucose-6-phosphate dehydrogenase enzyme. It is also the first committed enzyme in the pathway and is involved in the regulation. PPP metabolizes from 5 to 30% of glucose depending on the tissue type. The enzymatic reaction is: 1. Glucose-6-phosphate dehydrogenase: Glc6P + NADP+ → 6-phosphogluconolactone + NADPH The oxidative branch produces NADPH for reductive biosynthesis and it also maintains the reducing condition of the cell to protect it against oxidative stress, which is especially important in erythrocytes (Figure\(3\)). It also starts the pathway to produce 5-carbon sugars for nucleotide biosynthesis. G6PDH and ROS G6PDH is very important for the protection of free radicals in red blood cells since they don't have mitochondria that could provide another source of NADPH for protection. NADPH is involved in protection against ROS through the three cycles shown in Figure \(3\). Figure \(3\): Function of G6PD enzyme in the PPP from red blood cells. In G6PD-normal red cells, the NADPH is produced by the action of glucose 6-phosphate dehydrogenase (G6PD) and 6-phosphogluconate dehydrogenase (6PGD) enzymes. The NADPH serves as a proton donor to regenerate the GSSG oxidized. Cat = Catalase; GPx = Glutathione peroxidase; GR = Glutathione reductase; G6PD = glucose 6-phosphate dehydrogenase; 6PGL = 6-phosphogluconolactonase; 6GPD = 6-phosphogluconate dehydrogenase; SOD = Superoxide dismutase; GSH = Reduced glutathione; GSSG = Oxidized glutathione; H2O2 = Peroxide; O2 = Superoxide. Gomez-Manzo et al. 2016 Dec; 17(12): 2069. doi: 10.3390/ijms17122069. Creative Commons Attribution (CC-BY) license (http://creativecommons.org/licenses/by/4.0/). NADPH can keep glutathione in its reduced form, which as a substrate for catalase can help rid the cell of hydrogen peroxide and indirectly other ROS. The enzyme is active as a dimer or tetramer. Each monomer in the complex has a substrate binding site that binds to G6P, and a catalytic coenzyme binding site that binds to NADP+/NADPH. For some higher organisms, such as humans, G6PD contains an additional NADP+ binding site, called the NADP+ structural site, that does not seem to participate directly in the reaction catalyzed by G6PD. The evolutionary purpose of the NADP+ structural site is unknown, however, it does play a role in the overall stability of the enzyme. Mutations that cause dysfunction or deficiencies are very common, especially in males and in Africa, Asia, the Mediterranean, and the Middle East, in a geographic distribution that parallels the incidence of malaria. Interestingly, many of the mutations occur near the NADP+ structural site. Glucose-6-phosphate dehydrogenase deficiency is very common worldwide and causes acute hemolytic anemia in the presence of simple infection, ingestion of fava beans, or reaction with certain medicines. An abbreviate mechanism for glucose-6-phosphate dehydrogenase from Leuconostoc mesenteroides is shown in Figure \(4\). Figure \(4\): Mechanism of Leuconostoc mesenteroides glucose-6-phosphate dehydrogenase. Ribeiro AJM et al. (2017), Nucleic Acids Res, 46, D618-D623. Mechanism and Catalytic Site Atlas (M-CSA): a database of enzyme reaction mechanisms and active sites. DOI:10.1093/nar/gkx1012. PMID:29106569 https://www.ebi.ac.uk/thornton-srv/m-csa/entry/843/. Creative Commons Attribution 4.0 International (CC BY 4.0) License. Figure \(5\) shows an interactive iCn3D model of the glucose 6-phosphate dehydrogenase from Leuconostoc mesenteroides (1DPG). Figure \(5\): Glucose 6-phosphate dehydrogenase from Leuconostoc mesenteroides (1DPG). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...m8NCXiBuQbTQb7 Interestingly, this bacteria has an incomplete glycolytic pathway and can use this enzyme with either NADP+ for anabolism or NAD+ for catabolism. Each monomer can bind two NADP+, one at a site that promotes structure integrity and stability and the other at the catalytic site where NADP+ serves as a substrate (or cofactor). The other substrate, glucose-6-phosphate bind between the two. Figure \(6\) shows an interactive iCn3D model of the human glucose 6-phosphate dehydrogenase with bound structural and substrate NADP+ (2BH9). Figure \(6\): Human glucose 6-phosphate dehydrogenase with bound structural and substrate NADP (2BH9).(Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?NFJnS7UtUt3Uwu2D7 The biological unit shown is a dimer. The structural NADP+ is shown in spacefill bound in the N-terminal end of each monomer, while the substrate (cofactor) NADP+ is shown in sticks. Arg 459 is shown in spacefill. A common disease-causing mutation of G6PDH is the Canton R459L mutation. It's common in China and Southeast Asia. The activity of the enzyme is significantly decreased and the enzyme is less able to form tetramers. It unfolds at a lower temperature. This suggests that the mutation causes a significant conformation change. The Arg459 is shown in the above model. Its contribution to interhelical noncovalent attractions between it and D181 and N185. D181 is shown in proximity to Arg459 in the above model. Figure \(7\)s shows a static image of glucose 6-phosphate dehydrogenase with bound structural and substrate NADP+ as well as glucose-6-phosphate. Figure \(7\): Glucose 6-phosphate dehydrogenase with bound structural and substrate NADP+ as well as glucose-6-phosphate (2BHL and 2BH9). Structural NADP+ (blue molecular surface), catalytic NADP+ (dark purple molecular surface), and G6P substrate (yellow molecular surface) in the dimer are shown. The two monomers are shown in cyan and green. Right inset, close-up of the dimer interface and both structural NADP+ molecules. Gomez-Manzo et al., ibid. The second reaction of the oxidative branch is mediated by the phosphogluconolactonase (6PGL, PGLS) enzyme. The overall reaction is shown here: 2. 6-phosphogluconolactamase: 6-phosphogluconolactone + H2O → 6-phosphogluconate 6PGL is a cytosolic enzyme found in all organisms that catalyzes the hydrolysis of 6-phosphogluconolactone to 6-phosphogluconic acid in the oxidative phase of the pentose phosphate pathway. 6PGL hydrolysis of 6-phosphogluconolactone to 6-phosphogluconic acid has been proposed to proceed via proton transfer to the O5 ring oxygen atom (Figure \(8\)). The reaction initiates via the attack of a hydroxide ion at the C5 ester. A tetrahedral intermediate forms and the elimination of the ester linkage follows, aided by the donation of a proton from an active site histidine residue. Figure \(8\): Mechanism for6-phosphogluconolactamaseHis 165 and Asp 163 appear to be involved in a proton relay scheme. His 163 is conserved in lactonase. Arg 77 and 200 are involved in binding the substrate. Figure \(9\) shows an interactive iCn3D model of the 6-phosphogluconolactonase from Trypanosoma brucei complexed with 6-phosphogluconic acid (3E7F). This enzyme is a target for developing drug treatment strategies for African sleeping sickness. Figure \(9\): 6-phosphogluconolactonase from Trypanosoma brucei complexed with 6-phosphogluconic acid (3E7F). The key residues involved in binding and the charge relay system are shown in sticks and labeled. The product, 6-phosphogluconic acid, is shown in spacefill. The single sphere is a Zn2+ which does not appear to be involved in catalysis. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?jRgi1nfrSGGgTYWo8 Glyoxosomes and glycosomes Trypanosomiasis is a disease usually referring to African human trypanosomiasis or African sleeping sickness. This infectious disease is caused by the parasites Trypanosoma brucei gambiense or Trypanosoma brucei rhodesiense cause this infectious disease, and the tsetse fly transmits the disease. The vector tsetse fly, Glossina, carries the trypanosome within the midgut after a blood meal. These protozoa then migrate to the salivary glands of the fly whereby they can be transmitted during the next feeding. After inoculation within the host, the parasite can live freely within the bloodstream and evade mammalian host defenses through variable surface glycoproteins (VSG). The clinical disease has 2 stages. These are characterized by an early/first hemolymphatic stage and a late /second meningoencephalitis stage with an invasion of the central nervous system (CNS). In stage 1, systemic symptoms develop including intermittent fever, headache, pruritus, and lymphadenopathy. Undulating fevers reflect parasites multiplying within the blood. Less frequent hepatosplenomegaly may occur in the early stage. In the late/second stage, CNS symptoms manifest as sleep disturbances or neuropsychiatric disorders. A sleep disorder is the most common symptom of the second stage, and it is from this that the term “African sleeping sickness” was ascribed. New Drug Targets: Enzymes involved in the pentose phosphate pathway provide a new set of drug targets to help combat parasitic infections such as those caused by trypanosomes. In this context, a focus for new therapeutics has been on the disruption of glycosomes within these organisms. Glyoxosomes are specialized peroxisomes found in plants and some fungi. They all have enzymes for the glyoxylate shunt and can hence create some glucogenic intermediates. In trypanosomes and Leishmania, another internal organelle called the glycosome is found. This houses most of the glycolytic enzymes and also the oxidative enzymes of the pentose phosphate pathway. This makes the enzymes in the glycosome unique drug targets. The third enzyme utilized in the oxidative branch of the PPP is 6-phosphogluconate dehydrogenase (6PGDH). The overall reaction is shown here: 3. 6-phosphogluconate dehydrogenase: 6-phosphogluconate + NADP+ ↔ ribulose-5-phosphate + NADPH + CO2 6PGDH catalyzes a reversible oxidative decarboxylation reaction, as shown in Figure \(10\). Oxidation, followed by decarboxylation forms an endiol intermediate followed by conversion to product. Notably, NADP+ serves as an electron acceptor in the reaction, leading to the production of a second molecule of NADPH. Essentially, an active center lysine abstracts a proton from the substrate, 6-phosphogluconate, and NADP+ is reduced, causing the formation of a ketone-intermediate. Decarboxylation leads to the formation of the enediol intermediate. The active site lysine abstracts a proton in the second half of the reaction causing the formation of ribulose 5-phosphate. The abstracted proton is transferred to the active site Glu190 resetting the enzyme for another catalytic cycle. Figure \(10\): Mechanism of 6-phosphogluconate dehydrogenase. Figure \(11\) shows an interactive iCn3D model of sheep 6-phosphogluconate dehydrogenase (2PGD). Figure \(11\): Sheep 6-phosphogluconate dehydrogenase (2PGD). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...DqosWmFg1DqK16 The protein is a homodimer in which the monomers act independently: each contains a large, mainly alpha-helical domain and a smaller beta-alpha-beta domain, containing a mixed parallel and anti-parallel 6-stranded beta-sheet. NADP+ is bound in a cleft in the small domain, the substrate binding in an adjacent pocket. Nonoxidative branch The non-oxidative branch of the pentose phosphate pathway (nonox-PPP) is responsible for the formation of simple sugars within the cell and maintaining these building blocks in appropriate concentrations. The enzymes utilized in this pathway mediate a set of reversible reactions that lead to the production of ribose-5-phosphate, xylulose-5-phosphate, sedoheptulose 7-phosphate, and erythrose 4-phosphate, as well as intermediates utilized in the glycolytic pathway, including glyceraldehyde-3-phosphate and fructose-6-phosphate Major enzymes utilized include transketolase (TKT) and transaldolase (TALDO), as well as important isomerase and epimerase enzymes. Ribose 5-phosphate is an important building block for the biosynthesis of nucleotides, and erythrose 4-phosphate is used in the synthesis of aromatic amino acids. When these simple sugars are in excess, they can also be converted into glycolytic intermediates and utilized for energy production. The nonoxidative branch of the phosphopentose pathway is shown in Figure \(12\). Figure \(12\) :Nonoxidative branch of the phosphopentose pathway Now let's examine the individual enzymes. Ribulose 5-phosphate is the starting place for the non-oxidative portion of the PPP. It can be converted down two different pathways, either to ribose-5-phosphate or to xylulose-5-phosphate. The ribose 5-phosphate isomerase is involved in the ketose-aldose conversion and will be the first enzyme discussed. Ribose-5-phosphate isomerase (Rpi) This enzyme is fully reversible and is usually referred to as ribose-5-phosphate isomerase, although it can also be known as ribulose 5-phosphate isomerase, as it mediates the transition between this aldose-ketose pair. In addition to the PPP, this enzymatic reaction is also required in the Calvin cycle in photosynthesis. There are two different Rpi enzymes, RpiA and RpiB, which have little sequence or structural similarities as well as different mechanisms. RpiA is found in all three kingdoms of life and is highly conserved due to its role in the PPP and the Calvin Cycle of photosynthesis. RpiB, on the other hand, is only found in some bacteria and protozoans. Thus, RpiB is a potential therapeutic target for the treatment of diseases such as African Sleeping Sickness, Chagas disease, and leishmaniasis. While RpiA and RpiB are structurally very different, both enzymes catalyze the isomerization reaction through an enediol intermediate using the linear form of the sugar. The reaction mechanism of RpiB is detailed below. Figure \(13\) shows a mechanism for Escherichia coli RpiB. Figure \(13\): Mechanism for Escherichia coli RpiB. https://www.ebi.ac.uk/thornton-srv/m-csa/entry/680/. Creative Commons Attribution 4.0 International (CC BY 4.0) License. RpiB is a homodimer that can form a tetramer. The monomer subunits are coordinated in a head-to-tail format, such that there are two active sites per dimer where side chains from each monomer contribute to each active site (Figure \(14\)). The reaction mechanism requires two major steps: (1) ring opening of the furanose form of the sugar, and (2) isomerization through an enediol intermediate (Figure \(13\)). Figure \(14\) shows an interactive iCn3D model of Ribose-5-phosphate isomerase (RPIB_AlsB) from Escherichia coli (1NN4). Figure \(14\): Ribose-5-phosphate isomerase (RPIB_AlsB) from Escherichia coli (1NN4).Active site residues are indicated in each of the monomer subunits. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...ByuLUqhhs9B8c9 The other major pathway for ribulose 5-phosphate conversion involves the ribulose 5-phosphate (3) epimerase (RPE) and the conversion to xylulose 5 phosphate. Ribulose-5-phosphate (3) epimerase (RPE) This enzyme is also called pentose-5-phosphate 3-epimerase or PPE. Sugar epimerase enzymes alter the stereochemistry of the sugar at one of the alcohol positions creating a different epimer. The RPE enzyme has a Zn2+ cofactor, however, its requirement for enzyme activity is not essential. A Zn2+ independent form can still function and stabilize an oxyanion intermediate with adjacent and conserved methionines. A mechanism of the Zn2+ dependent form of the enzyme is shown in Figure \(15\). Figure \(15\): Mechanism for rice ribulose-5-phosphate epimerase The enzyme utilizes an acid/base catalytic mechanism that mediates the formation of a trans-2,3-enediol phosphate intermediate. Key aspartic acid residues act as proton donors and acceptors during the reaction. A zinc metal cofactor helps stabilize charges during the reaction. Figure \(16\) shows an interactive iCn3D model of cytosolic D-ribulose-5-phosphate 3-epimerase from rice (1h1z). Figure \(16\): Cytosolic D-ribulose-5-phosphate 3-epimerase from rice (1h1z). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...AH7uS2T8hsnpXA Transketolase (TK) The pool of ribose 5-phosphate that is created can be utilized for nucleotide production, or some of it can undergo a reaction with the xylulose 5-phosphate created in the RPE reaction. The transketolase enzyme converts two five-carbon sugars (ribose 5-phosphate and xylulose 5-phosphate) into a 3-carbon and a 7-carbon sugar (glyceraldehyde 3-phosphate and sedoheptulose 7-phosphate). Essentially, transketolase enzymes transfer ketone functional groups from ketoses to aldoses, effectively creating a new ketose that is two carbons larger. The ketose donor then becomes an aldose with two fewer carbons. The basic transketolase reaction shown in Figure \(17\) Figure \(17\): Transketolase reaction. The reaction shows the reversible conversion of ribose 5-phosphate and xylulose 5-phosphate to sedoheptulose 7-phosphate and glyceraldehyde 3-phosphate. Note the number of carbons in the reactants and products: 5C + 5C ↔ 3C + 7C. In this reversible reaction, the enzyme uses the cofactor thiamine pyrophosphate (TPP) and a divalent cation. The enzyme transfers a 2C ketol group from xylulose-5-phosphate to ribose-5-phosphate, an aldose. The product glyceraldehyde-3-phosphate is a glycolytic intermediate and can be used in the glycolytic pathway. The mechanism for the reverse reaction (yeast numbering system) is shown in Figure \(18\). In this reaction, sedoheptulose 7-phosphate binds with the enzyme and the TPP cofactor is activated to form a carbanion. The carbanion mediates nucleophilic attack on the carbonyl carbon of the substrate forming a covalent intermediate. His263 serves as a base and abstracts a proton, enabling bond cleavage and the formation of ribose 5-phosphate. Ribose 5-phosphate leaves the active site and glyceraldehyde 3-phosphate enters. The two carbon intermediate covalently bound to the TPP mediates nucleophilic attack on the glyceraldehyde 3-phosphate enabling the formation of xylulose 5-phosphate and the restoration of the enzyme. Figure \(18\): Mechanism of the transketolase reaction during the pentose phosphate pathway. https://www.ebi.ac.uk/thornton-srv/m-csa/entry/219/ Figure \(19\) shows an interactive iCn3D model of Human transketolase in a covalent complex with donor ketose D-xylulose-5-phosphate (4kxv). Figure \(19\): Human transketolase in covalent complex with donor ketose D-xylulose-5-phosphate (4kxv). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...WtGtQ7mKA6UYx7 The enzyme is a dimer (gray and cyan coloring). Numbering of key residues compared to yeast mechanism: Human Yeast E366 E418 Q428 H481 H258 H262 H38 H30 Also shown are human active site residues H474, R318, and S345. The covalent xylulose-5-phosphate adduct is shown in spacefill and labeled DX5 (xylitol-5-phosphate). TPP is shown in sticks and labeled. The metal ion is Ca2+. Structural analyzes show a 200 distortion in the planarity of the cofactor-substrate bond and a lengthening of the C-C bond in the substrate which breaks. Transaldolase In addition to being used as a resource in the glycolytic pathway, glyceraldehyde 3-phosphate can also be utilized as a substrate in a transaldolase reaction along with the sedoheptulose 7-phosphate that is produced in the previous reaction. This results in the formation of erythrose 4-phosphate and fructose 6-phosphate. A summary of the reaction is shown in Figure \(20\). Figure \(20\): Summary of transaldolase reaction As with transketolase, the transaldolase enzyme is reversible. Again, note the number of carbons in the reactants and products: 3C + 7C ↔ 4C + 6C. Unlike the transketolase used in the last reaction, the transaldolase enzyme does NOT use TPP as a cofactor. Instead, it forms a Schiff base intermediate similar to that of the aldolase enzyme in the glycolytic pathway. The enzyme removes a 3C ketol group (dihydroxyacetone) from sedoheptulose 7-phosphate and transfers it to glyceraldehyde 3-phosphate forming fructose 6-phosphate. Erythrose 4-phosphate is left from the original sedoheptulose 7-phosphate. We will explore the mechanism for the E. Coli enzyme in the reverse direction. The first part of the mechanism of transaldolase is shown in Figure \(21\). Figure \(21\): Mechanism for the first half of the transaldolase reaction. https://www.ebi.ac.uk/thornton-srv/m-csa/entry/148/ In the first part of the reaction, fructose 6-phosphate binds to the active site of the enzyme, where an active site lysine residue mediates nucleophilic attack on the carbonyl carbon and forms a covalent intermediate with the enzyme. Formation of the Schiff base leads to dehydration of the intermediate. The Schiff base nitrogen becomes protonated and this leads to the oxidation of the C4 hydroxyl leads and subsequent bond cleavage releasing glyceraldehyde 3-phosphate. The remaining ES-complex rearranges to form an enol intermediate. The mechanism for the second half of the transaldolase reaction is shown in Figure \(22\). Figure \(22\): Mechanism for the second half of the transaldolase reaction. https://www.ebi.ac.uk/thornton-srv/m-csa/entry/148/ Once glyceraldehyde 3-phosphate has left the active site of the enzyme, erythrose 4-phosphate can bind. The enol from the ES intermediate mediates nucleophilic attack on the aldehyde carbonyl group of erythrose 4-phosphate. This results in the formation of a Schiff base intermediate. Hydration at the Schiff base carbon atom ensues followed by the oxidation of the newly incorporated alcohol to form a ketone functional group. Formation of the ketone causes bond cleavage between the enzyme and the newly formed ketose, sedoheptulose 7-phosphate. Figure \(23\) shows an interactive iCn3D model of transaldolase B from Escherichia coli (1ONR). Figure \(23\): 3D structure of Transaldolase B from E. coli (1ONR).  (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...7swLZVpNPe3eU9 Transketolase As seen above, fructose 6-phosphate is a product of the transaldolase enzyme during the nonoxidative branch of the PPP. This sugar can feed back into the glycolytic cycle. The formation of fructose 6-phosphate can also occur using the transketolase enzyme when it transfers a 2C ketol group from xylulose 5-phosphate to the aldose, erythrose 4-phosphate. This reaction is summarized in Figure \(24\). Figure \(24\) Summary of second transketolase reaction during the pentose phosphate pathway. Both products of this final transketolase reaction can be utilized in energy formation through glycolysis. Thus, there are clear metabolic ties of the PPP with the energy-producing glycolytic pathway and intermediates from both pathways can easily be exchanged through the activity of these reversible enzymes. The PPP provides key intermediates, including ribose 5-phosphate that is used in the biosynthesis of nucleotides and ultimately nucleic acids, as well as, other important metabolic molecules such as FAD, NAD+, NADP+, and Coenzyme A. In addition to being a biosynthetic source for these molecules, the PPP also is the primary source for reducing NADP+ to NADPH, during the oxidative reactions. NADPH is utilized in many biosynthetic processes as an electron donor. For example, the biosynthesis of many lipid molecules requires NADPH, including the production of triacylglycerols, phospholipids, and steroids. NADPH is also required for the biosynthesis of some amino acids (such as glutamate and proline) and is also involved in the reduction of ribonucleotides and deoxyribonucleotides during the synthesis of RNA and DNA. NADPH is also utilized by a number of oxidoreductases involved in detoxification reactions within the body. Within the immune system, NADPH oxidases or NOX enzymes are involved in the production of superoxide and utilized to damage invading pathogens nonspecifically. NADP+/NADPH ratios also can play a regulatory role in cellular metabolic processes and are utilized as allosteric effectors for several enzymes and oxidation sensor proteins. Regulation of the Pentose Phosphate Pathway The cellular demand for the two major products of the PPP (ribose 5-phosphate and reduced NADPH) can be different depending on the cell type or the current cellular environment, such as times of increased metabolic demand or oxidative stress. Thus, the two major products may need to be produced in different quantities and independently of one another. For example, we can imagine times when the needs for ribose 5-phosphate and NADPH are in balance with the standard PPP reactions. However, we can also imagine times when the demand for ribose 5-phosphate may be much higher than the demand for NADPH, or vice versa, the demand for NADPH may be much greater than the demand for ribose 5-phosphate. Thus, there are regulatory strategies in place that enable the ability to regulate the production of these different pools independently of one another and adapt to cellular needs. This ability is largely dependent on the production of metabolic intermediates that can be easily interchanged within the glycolytic pathway. For example, the ratio of NADP+/NADPH serves as a key regulator of the oxidative branch of the PPP. The first enzymatic step of the pathway mediated by the glucose 6-phosphate dehydrogenase (G6PD) is regulated in this fashion and helps control the pool of glucose 6-phosphate that will be utilized within the PPP to produce reduced NADPH. Low levels of NADP+ inhibit the G6PD enzyme. The G6PD reaction is essentially irreversible and serves as the committed step for glucose to enter into the oxidative portion of the PPP. Thus the regulation of this enzymatic step is key in the regulation of NADPH levels within the cell. When the ratio of NADP+/NADPH increases, G6PD becomes more active and the reduction of NADP+ to NADPH increases. When ribose 5-phosphate is also in high demand, the pool of ribose 5-phosphate will be low and increase the activity of the ribose 5-phosphate isomerase in the forward direction to convert the ribulose 5-phosphate produced in the oxidative branch to generate more ribose 5-phosphate. However, if more ribose 5-phosphate is required than can be supplied from the activity of the oxidative branch, the pool of ribose 5-phosphate can be fed from the conversion of intermediates from the glycolytic pathway in the nonoxidative branch of the PPP. The opposite occurs as well when there is a high need for NADPH in the cell but a low need for ribose 5-phosphate. In this situation, ribose 5-phosphate is converted to fructose and glyceraldehyde 3-phosphate and can be incorporated into the glycolytic pathway. In addition, the activity of the PPP varies depending on the tissue type and location within the body. For example, skeletal muscle has very low PPP activity, as this tissue requires more energy production and activity of the glycolytic pathway. On the other hand, the PPP is highly active in adipose tissue due to the heightened requirement for intermediates needed for lipid biosynthesis.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/13%3A_Glycolysis_Gluconeogenesis_and_the_Pentose_Phosphate_Pathway/13.04%3A_Pentose_Phosphate_Pathway_of_Glucose_O.txt

• 14.1: Regulation of Metabolic Pathways Exquisite mechanisms have evolved that control the flux of metabolites through metabolic pathways to insure that the output of the pathways meets biological demand and that energy in the form of ATP is not wasted by having opposing pathways run concomitantly in the same cell. • 14.2: Basic Principles of Metabolic Control Analysis (MCA) Enzyme kinetics may seem difficult given the complicated mathematical derivations, the number of chemical species involved (an enzyme and all its substrates and products), the number of steps in the mechanism, and the large number of rate, kinetic, and dissociation constants. • 14.3: The Flux Control Coefficient Metabolic control analysis is one way to address the complexity of dynamic changes of species in a complex metabolic system. As such an understanding of MCA would apply to complex signal transduction pathways as well as to an understanding of the emerging discipline of systems biology. In MCA external inputs (source) and outputs (exits) pools or reservoirs exists which are connected to the internal metabolic enzymes, reactants and products of the pathway connecting the two external reservoirs. • 14.4: Concentration Control and Elasticity Coefficients • 14.5: Metabolism and Signaling: The Steady State, Adaptation and Homeostasis Thumbnail: Overview of regulatory interactions involved in metabolic regulatory networks. The function of metabolic networks are governed by constraints. The regulation of a metabolic network involves a tight interplay between different cellular networks such as signalling and gene networks and by interactions with its environment. The enzyme capacity is the net result of the amount of enzyme expressed and its activity as dictated by post-translational modification and allosteric regulation. Metabolite pools and fluxes are considered as the outputs of metabolic reaction networks and can be involved in various regulatory feedback loops to other networks within the metabolic reaction networks as indicated by the dashed arrows. (CC BY 3.0; Jan Berkhout, Frank J. Bruggeman, and Bas Teusink via MDPI) 14: Principles of Metabolic Regulation Search Fundamentals of Biochemistry Exquisite mechanisms have evolved that control the flux of metabolites through metabolic pathways to insure that the output of the pathways meets biological demand and that energy in the form of ATP is not wasted by having opposing pathways run concomitantly in the same cell. Enzymes can be regulated by changing the activity of a preexisting enzyme or changing the amount of an enzyme. Changing the activity of a pre-existing enzyme The quickest way to modulate the activity of an enzyme is to alter the activity of an enzyme that already exists in the cell. The list below, illustrated in the following figure, gives common ways to regulate enzyme activity 1. Substrate availability: Substrates (reactants) bind to enzymes with a characteristic affinity (characterized by a dissociation constant) and a kinetic parameter called Km (units of molarity). If the actual concentration of a substrate in a cell is much less than the Km, the activity of the enzyme is very low. If the substrate concentration is much greater than Km, the enzyme's active site is saturated with substrate and the enzyme is maximally active. 2. Product inhibition: A product of an enzyme-catalyzed reaction often resembles a starting reactant, so it should be clear that the product should also bind to the activity site, albeit probably with lower affinity. Under conditions in which the product of a reaction is present in high concentration, it would be energetically advantageous to the cell if no more product was synthesized. Product inhibition is hence commonly observed. Likewise, it is energetically advantageous to a cell if the end product of an entire pathway could likewise bind to the initial enzyme in the pathways and inhibit it, allowing the whole pathway to be inhibited. This type of feedback inhibition is commonly observed. Figure \(1\) shows product and end-product inhibition. Figure \(1\): Product and end product inhibition of an enzyme 1. Allosteric regulation: As many pathways are interconnected, it would be optimal if the molecules of one pathway affected the activity of enzymes in another interconnected pathway, even if the molecules in the first pathway are structurally dissimilar to reactants or products in a second pathway. Molecules that bind to sites on target enzymes other than the active site (allosteric sites) can regulate the activity of the target enzyme. These molecules can be structurally dissimilar to those that bind at the active site. They do so by conformational changes which can either activate or inhibit the target enzyme's activity. 2. pH and enzyme conformation: Changes in pH that can accompany metabolic processes such as respiration (aerobic glycolysis for example) can alter the conformation of an enzyme and hence enzyme activity. The initial changes are covalent (change in the protonation state of the protein) which can lead to an alteration in the delicate balance of forces that affect protein structure. 3. pH and active site protonation state: Changes in pH can affect the protonation state of key amino acid side chains in the active site of proteins without affecting the local or global conformation of the protein. Catalysis may be affected if the mechanism of catalysis involves an active site nucleophile (for example), that must be deprotonated for activity. 4. Covalent modification: Many if not most proteins are subjected to post-translational modifications which can affect enzyme activity through local or global shape changes, by promoting or inhibiting binding interaction of substrates and allosteric regulators, and even by changing the location of the protein within the cell. Proteins may be phosphorylated, acetylated, methylated, sulfated, glycosylated, amidated, hydroxylated, prenylated, or myristoylated, often in a reversible fashion. Some of these modifications are reversible. Regulation by phosphorylation through the action of kinases, and dephosphorylations by phosphates are extremely common. Control of the phosphorylation state is mediated through signal transduction processes starting at the cell membrane, leading to the activation or inhibition of protein kinases and phosphatases within the cell. Figure \(2\) shows ways to regulate the activity of pre-existing enzymes. Figure \(2\): Ways to regulate the activity of pre-existing enzymes Extracellular regulated kinase 2 (ERK2), also known as mitogen-activated protein kinase 2 (MAPK2) is a protein that plays a vital role in cell signaling across the cell membrane. Phosphorylation of ERK2 on Threonine 183 (Thr153) and Tyrosine 185 (Tyr185) leads to a structural change in the protein and the regulation of its activity. Figure \(3\) shows an interactive iCn3D model showing the structural alignment of ERK2 in the dephosphorylated (5UMO) and phosphorylated (pY185) forms (2ERK). Toggle back and forth between the two structures with the "a" key. The residues that change significantly in conformation on phosphorylation are shown in blue. The side chain of tyrosine 185 in the unphosphorylated form is shown in CPK-colored sticks and labeled. Regulation of single enzymes or entire pathways: Enzyme condensates Single enzymes or all the enzymes of a given pathway can be coordinately regulated to maximize end-product output by organizing the enzymes in one large complex built from soluble enzymes that produce a "condensate" through a process similar to phase separation. Such condensates are shown for a series of enzymes in Figure \(4\). Figure \(4\): Supramolecular assembly of enzyme condensates. Prouteau and Loewith. Biomolecules 2018, 8(4), 160; https://doi.org/10.3390/biom8040160. Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/). The figure shows metabolism-related enzymes that form polymers in various organisms. Panel (a) shows examples of metabolic enzymes observed to coalesce into cytosolic condensates. Panel (b) top shows structures of metabolic enzymes that polymerize into filaments. The protomer of the polymer is shown above and placed into the filament below. These include P-Fructo-Kinase (4XYJ), cytidine triphosphate synthase (5U03), acetyl-CoA carboxylase (6G2D), glutamine synthetase (3FKY), mTORC1 (5FLC) from PDB files. Panel (b) bottom shows the same structures from the Electron Microscopy Data Bank, including P-Fructo-Kinase filament (emd-8542), human CTP synthase filament (and-8474), human acetyl-CoA carboxylase with citrate (emd-4342), and the yeast glutamine synthetase filament. Changing the amount of an enzyme Another longer-duration method to modulate the activity of an enzyme is to alter the activity of an enzyme that already exists in the cell. Figure \(5\) shows ways in which enzyme concentrations are regulated. Methods include: 1. Alterations in the transcription of enzyme's gene: Extracellular signals (hormones, neurotransmitters, etc) can lead to signal transduction responses and ultimate activation or inhibition of the transcription of the gene for a protein enzyme. These changes result from the recruitment of transcription factors (proteins) to DNA sequences that regulate the transcription of the enzyme gene. 2. Degradation of messenger RNA for the enzyme: The levels of messenger RNA for a protein will directly determine the amount of that protein synthesized. Small inhibitor RNAs, derived from microRNA molecules transcribed from cellular DNA, can bind to specific sequences in the mRNA of a target enzyme. The resulting double-stranded RNA complex recruits an enzyme (Dicer) that cleaves the complex with the effect of decreasing the translation of the protein enzyme from its mRNA. 3. Co/Post-translational changes: Once a protein enzyme is translated from its mRNA, it can undergo many changes that regulate its activity. Some proteins are synthesized in a "pre" form which must be cleaved in a targeted and limited fashion by proteases to activate the protein enzyme. Some proteins are not fully folded and must bind to other factors in the cell to adopt a catalytically active form. Finally, fully active protein can be fully proteolyzed by the proteasome, a complex within cells, or in lysosomes, which are organelles within cells containing proteolytic enzymes. All proteins are ultimately regulated, if only by modulating the rates of their synthesis and degradation. However, some enzymes positioned at key points in metabolic pathways are ideal candidates for regulation, as their activity can affect the output of entire pathways. These enzymes typically have two common characteristics, they catalyze reactions far from equilibrium and they catalyze early committed steps in pathways. Which Enzymes to Regulate: Reactions not at Equilibrium The optimal enzymes for regulation are those at the beginning of pathways that carry out thermodynamically favored reactions. Why is the latter so important? These enzymes control the flux of metabolites through pathways, so to understand their regulation we can use the analogy of flow (or flux) of water from one container to another as illustrated in Figure \(6\). Figure \(6\): Regulation of water flow in pipe Let's say you wish to fill a swimming pool at any desired height you wish and you have two ways to do so (see figure below). You could open a valve that controls the flow from your town's water tower to the pool. In this, the reaction (flow of water) is energetically (thermodynamically) favored given the difference in height of the water levels and the potential energy difference between the two. Even though flow (or flux) is cleared flavored, you can regulate it, from no flow to maximal flow, by opening and closing the valve (analogous to activating and inhibiting an enzyme). Your choices in the other scenario, filling the pool from a lake, are not so great. It would be hard to fill the water to the desired level (especially if it was an above-ground pool). It would be hard to regulate the flow. By analogy, the best candidates for regulation are those enzymes whose reactions are thermodynamically favored (not at equilibrium) but which can be controlled by the mechanisms discussed in the previous section. Which reactions are commonly not at equilibrium (i.e. ΔG<0 and usually also ΔG0 <0 if the ratio of products to reactants is not too high)? The answer is those that have reactants that are thermodynamically unstable compared to their reaction products. Several types of reactions often fit these criteria: Hydrolysis (or similar reactions) of anhydride or analogous motifs: The figure below shows molecules with similar "anhydride" motifs and the ΔG0 for hydrolysis of the molecules. Those with more negative ΔG0 values can transfer their phosphate group to ADP to make ATP, which is necessary to drive unfavorable biological reactions. Metabolic reactions that involve hydrolysis (or other types of transfer reaction of these groups) usually proceed with a negative ΔG0 and ΔG, making them prime candidates for pathway regulation. Many textbooks label these types of molecules as having "high energy" bonds. This is confusing to many students as bonds between atoms lower the energy compare to when the atoms are not bonded. It takes energy to break the "high" energy phosphoanhydride covalent bond.  What make hydrolysis of the molecules below so exergonic is that more energy is released on bond formation within the new products than was required to break the bonds in the reactants. In addition, other effects such as preferential hydration of the products, lower charge density in the products, and less competing resonances in the products all contribute to the thermodynamically favorable hydrolysis of the reactants. Figure \(7\) shows thermodynamically unstable molecules (compared to their reaction products in aqueous solutions). Thioesters (such as Acetyl-SCoA) are also included as they have the same negative ΔG0 of hydrolysis as ATP, even though they lack an "anhydride" motif. Thioesters are destabilized compared to their hydrolysis products and in comparison to esters made with alcohol since the C-S bond is weaker. Why? Redox reactions: Everyone knows that redox reactions are thermodynamically favored if the oxidizing agent deployed is strong enough. The oxidation reactions of hydrocarbons, sugars, and fats by dioxygen are clearly exergonic (we do call these combustion reactions after all). What about redox reactions with less powerful oxidants? NAD+ is used frequently as a biological oxidizing agent. Are all these reactions as favored as combustion? Hardly so. Remember that in every redox reaction, an oxidizing and reducing agent react to form another oxidizing and reducing agent. Consider the following reaction: Pyruvate + NADH ↔ Lactate + NAD+. This reaction can go either way and is reversible. The above form is written in the favored direction in anaerobic metabolism when both Pyr and NADH levels are high. Although the ΔG0 favors the oxidation of lactate, given the high concentration of Pyr and NADH, the reaction is driven in the opposite direction and proceeds as shown. To determine if a redox reaction is favored and likely to occur (and possibly be regulated), the ΔG0 for a redox reaction should be calculated from standard reduction potentials, using the formula ΔG0 = -nFE0. Which Enzymes to Regulate: Those catalyzing committed steps in pathways The best enzymes to regulate are those that catalyze the first committed step in the reaction pathway. The committed step proceeds with a ΔG0 < 0 and is essentially irreversible. These reactions often occur from key metabolic intermediates that are immediately before or proximal to branches in reaction pathways. Two examples of key intermediates at branch points of metabolic pathways are shown in Figure \(8\) shows the reactions for the production and use of the intermediate glucose-6-phosphate. Figure \(10\) shows reactions for the production and use of the intermediate acetyl-CoA. Figure \(10\): Reactions for the production and use of the intermediate acetyl-CoA In reality, metabolic regulation is more complex and is distributed to many steps in a reaction pathway in ways that might not be evident without details mathematical analyses. We will discuss that in the next sections on metabolic control analysis.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/14%3A_Principles_of_Metabolic_Regulation/14.01%3A_Regulation_of_Metabolic_Pathways.txt

Search Fundamentals of Biochemistry Introduction to Metabolic Control Analysis Enzyme kinetics may seem difficult given the complicated mathematical derivations, the number of chemical species involved (an enzyme and all its substrates and products), the number of steps in the mechanism, and a large number of rate, kinetic, and dissociation constants. An example of such a “complicated” reaction explored earlier is shown in Figure $1$. v=\frac{\frac{k_2 k_3}{k_2+k_3}\left(E_0\right)(S)}{\left[\frac{k_3}{k_2+k_3}\right] K_S+S}=\frac{k_{c a t}\left(E_0\right)(S)}{K_M+S} But single enzymes rarely act in isolation. They are components of complex pathways which have a multitude of steps, many of which are regulated. To fully understand a reaction, it is important to study the concentrations of all species in the entire pathway as a function of time. Imagine deriving the equations and determining all the relevant concentrations and constants of a pathway such as glycolysis! To study enzyme kinetics in the lab, you have to spend much time developing assays to measure how the concentration of species changes as a function of time to be able to measure the initial velocities of an enzyme-catalyzed reaction. However, in networks of connected metabolic reactions, the concentration of some species in the system may not change. How can this happen? Two simple examples might help explain how. Example 1: There is no change in input or output from a given reaction. This would occur in a closed system for a reversible reaction at equilibrium. For a reversible reaction of reactant R going to product P (R ↔ P) with forward and reverse first-order rate constants, the following equation can be written at equilibrium: \begin{gathered} v_f=k_1 R=k_2 P \ K_{e q}=\frac{P_{e q}}{R_{e q}}=\frac{k_1}{k_2} \end{gathered} At equilibrium, $R$ and $P$ don’t change. Example 2: Consider the reaction as part of a pathway of reactions (like an open system). Now imagine a nonzero input to form reactant R and a nonzero output that consumes product P as shown in Figure $2$. If the input and output rates are the same, the concentrations of R and P would not change with time. That is the rate of formation of a reactant R for a given reaction is equal to the rate at which the product P of the given reaction is used. This would lead to a steady state but not equilibrium concentrations of the species. The Steady State - A Threshold Biochemistry Concept Students have a difficult time understanding the steady state and recognizing when it may prevail under a set of conditions. in vitro vs. in vivo Cornish-Bowden in his book Fundamentals of Enzyme Kinetics talks about key differences between the enzyme kinetics we do in test tubes (in vitro) and what happens in cells (in vivo). We've discussed this previously but it is important to reconsider it now. The conditions under which the enzymes are studied (in vitro) and operate (in vivo) are very different. • In vitro (in the lab), the enzyme is held at a constant concentration while the substrate is varied (i.e the substrate concentration is the independent variable). The velocity is determined by the substrate concentration. When inhibition is studied, the substrate is varied while the inhibitor is held constant at several different fixed concentrations. • In vivo (in the cell), the velocity might be held at a relatively fixed level in a pathway with the substrate determined by the velocity. To avoid a bottleneck in flux, substrate can't build up at the enzyme, so the enzyme processes it in a steady-state fashion to produce the product as determined by the Michael-Menten equation These differences are vastly underappreciated and not understood by students and instructors alike. Additionally, we confound our efforts in helping students understand the steady state when almost all of our efforts are focused on presenting initial rate v0 vs [S] curves when the substrate concentration is changed. Then instructors expect students to magically understand the steady state when substrate levels in pathways don’t change. It’s a big leap out of this box. We can help students better understand the steady state by shifting to progress curves (concentrations vs time), which are easily constructed by using programs such as VCell and Copasi. Let's use Vcell to analyze a very simple reversible enzyme in isolation, S ↔ P. Then we will place that same enzyme into a "mini" pathway, A ↔ S ↔ P ↔ Q in which there is one preceding reactant A and one following product Q that help control S and P levels. We have to start the simulation at some specified concentration so the default for the simulations is set such that [S] and [A] at time t = 0, S0 and A0, are 5, and the rest are set at 0. Hence as the simulation starts, there will be a readjustment of concentration until equilibrium or steady state concentration is reached. The reaction diagrams and their parameters are shown in Figure $3$. A reversible enzyme in isolation, S ↔ P A reversible enzyme for S ↔ P in a mini-pathway A ↔ S ↔ P ↔ Q Figure $3$: Reactions schemes and parameters for simple enzyme-catalyzed reversible reaction and the same reaction embedded in a "mini" pathway. Vcell simulation for the isolated enzyme Remember that an enzyme does not change the thermodynamics of a given reaction and hence doesn't alter Keq. It just speeds up both the forward and the reverse reactions. Hence you can calculate the Keq for the reaction condition from the Vcell model time course graph: \mathrm{K}_{\mathrm{eq}}=\frac{[\mathrm{P}]_{\mathrm{eq}}}{[\mathrm{S}]_{\mathrm{eq}}}=\frac{4}{1}=1 Now you observe that in this reaction, S does change from its initial value, S0 = 5 (since P0 = 0). Soon, however, the reaction comes to a real dynamic equilibrium in that both S and P don't change with time. You could choose a different initial concentration of S and P, rerun the simulation and calculate KEQ for the new conditions using the csv-downloaded spreadsheet data fro each run you make. They should be the same. MODEL Reversible reaction E + S ↔ ES  ↔  EP  ↔  E + P Initial values Select Load [model name] below Select Start to begin the simulation. Interactive Element Select Plot to change Y axis min/max, then Reset and Play  |  Select Slider to change which constants are displayed |  Select About  for software information. Move the sliders to change the constants and see changes in the displayed graph in real-time. Time course model made using Virtual Cell (Vcell), The Center for Cell Analysis & Modeling, at UConn Health.  Funded by NIH/NIGMS (R24 GM137787); Web simulation software (miniSidewinder) from Bartholomew Jardine and Herbert M. Sauro, University of Washington.  Funded by NIH/NIGMS (RO1-GM123032-04) Vcell simulation for the the enzyme in a "mini-pathway" Now compare this same reaction but in which S and P are part of the "mini-pathway" A ↔ S ↔ P ↔ Q, as shown in the Vcell model below. The enzyme kinetic parameters for S ↔ P (KM forward, VM forward, KM reverse, VM reverse) in the A ↔ S ↔ P ↔ O pathway were made the same as for the simple S ↔ P reaction since its the same enzyme. How do the A ↔ S and P ↔ Q reactions affect the apparent KEQ? Run the Vcell model with the defaults automatically set to the values in the table above! MODEL Initial Values Select Load [model name] below Select Start to begin the simulation. Interactive Element Select Plot to change Y axis min/max, then Reset and Play  |  Select Slider to change which constants are displayed |  Select About  for software information. Move the sliders to change the constants and see changes in the displayed graph in real-time. Time course model made using Virtual Cell (Vcell), The Center for Cell Analysis & Modeling, at UConn Health.  Funded by NIH/NIGMS (R24 GM137787); Web simulation software (miniSidewinder) from Bartholomew Jardine and Herbert M. Sauro, University of Washington.  Funded by NIH/NIGMS (RO1-GM123032-04) As in the first simulation, constant values for S and P are soon reached. Both are significantly lower than in the simple reaction of S ↔ P since P is being pulled toward Q faster than Q is converted back to S. Note also that Q reaches a higher concentration than either A or S but remember that the sum of the initial values of A and S is 10. Run the simulation again only this time set kr=2000 for the P ↔ Q reaction. Now calculate the "KEQ apparent" for the different kf values of the conversion of P → Q from this equation.  Again use the csv-downloaded spreadsheet data for each run you make. K_{\text {eq apparent }}=\frac{[\mathrm{P}]_{\text {steady state }}}{[\mathrm{S}]_{\text {steady state }}} Do they equal 4 as in case 1? No, they do not. You should see that the "KEQ apparent" value deviates most from 4 (lower number) when the rate constant for removal of P is highest (kf=2000). Animations Now let's look at some animation for these reactions as an additional way to understand the dynamics of the reactions. Click the image in Figure $4$ to view (in a new window) the animation of the chemical species and an inserted graph showing concentration vs time for the reversible enzyme-Catalyzed Reaction S ↔ P Pay attention to the disappearance of S (red) and the appearance of P (blue) species after interacting with the enzyme (green). Now here are two animations for the S ↔ P reaction when its embedded in the "minipathway" A ↔ S ↔ P ↔ Q when kf for the reaction P ↔ Q is 20 (left) and 200 (right). Click the images in Figure $5$ to view (in a new window) an animation of the chemical species and inserted graphs showing concentrations vs time. "mini-pathway" A ↔ S ↔ P ↔ Q, kf for the reaction P ↔ Q = 20 "mini-pathway" A ↔ S ↔ P ↔ Q, kf for the reaction P ↔ Q = 200 Figure $5$: Animation of the reversible enzyme-Catalyzed Reaction embedded in a "minipathway" A ↔ S ↔ P ↔ Q when kf for the reaction P ↔ Q is 200. These animations should reinforce your understanding of the differences between equilibrium and steady-state conditions, although you can calculate the actual KEQ apparent from the insert graphs without numerical data. Understanding a pure enzyme in vitro and in vivo requires different approaches. Biochemists like to isolate and purify to homogeneity an enzyme found in some tissue and study its mechanism of action. In doing thermodynamic measurements to measure equilibrium constants (Keq) or dissociation constants (KD), from which ΔG0 can be calculated, a protein concentration is usually held constant as the binding ligand concentration is varied (independent variable). A dependent variable signal (often spectroscopic) is measured. Measurements are made when equilibrium is reached. For enzyme kinetic measurements in vitro, the enzyme concentration is usually held constant while substrate and modifiers are varied (independent variables) to determine how velocity (dependent variable) changes. The velocity is determined by the substrate concentration. When inhibition is studied, the substrate is varied while the inhibitor is held constant at several different fixed concentrations. In vivo, the substrate concentration and even the enzyme concentration are determined by the velocity. Again compare this to in vitro kinetics when concentrations determine the velocity. For sets of reactions in pathways, it is better to use the term flux, J. In the steady state, the in and out fluxes for a given reaction are identical. Flux J is used to describe the rate of the system whereas rate or velocity v is used to describe the rate of an individual enzyme in a system. Computer programs can find steady-state concentrations by finding the roots of the ordinary differential equations (ODE) when set to zero (vf = vr). To model a process at very low concentrations, programs can also use probabilistic or stochastic simulations to model probability distributions for species and their change with time for a finite number of particles. In such simulations, concentrations (mM) are placed with the number of particles. ODEs don’t work well to describe these conditions since changes in concentrations are not continuous. Now back to our earlier rhetorical question of deriving the equations and determining all the relevant concentrations and constants of a pathway such as glycolysis! It has been done by Teusink et al for glycolysis in yeast. Many such complicated metabolic and signal transduction pathways have been mathematically modeled in the hopes of better understanding cellular and organismal responses. Quantitatively modeling and predicting input, outputs, and concentrations of all species in complex pathways is the basis of systems biology. Basic Principles of Metabolic Control Analysis - Glycolysis Let's use a more complex example, glycolysis, to illustrate the powers of computational modeling of entire pathways.  The reference for this model is shown below. Can yeast glycolysis be understood in terms of in vitro kinetics of the constituent enzymes? Testing biochemistry, Bas Teusink, Jutta Passarge, Corinne A. Reijenga, Eugenia Esgalhado, Coen C. van der Weijden, Mike Schepper, Michael C. Walsh, Barbara M. Bakker, Karel van Dam, Hans V. Westerhoff, and Jacky L. Snoep, 2000, European Journal of Biochemistry, 267, 5313-5329. PubMed ID: 10951190. Databases containing curated models with all the above information have been developed for many pathways. The yeast glycolysis model described in the reference above and the material below is found in the Biomodels Database. The model, BIOMD0000000064, can be downloaded as a systems biology markup language (SBML) file and imported into any of the programs described above. A variety of inputs are required for such computational analyses: a. Defined pathways. These are available in many databases. An example from the KEGG pathways for yeast glycolysis is shown in the left panel of Figure $6$ below. The right panel shows a more familiar representation, with glucose on the outside of the cell (Glco) entering the cell. b. A computational modeling program to input all parameters, equations, and models and calculate concentrations for all species as a function of time. We have been using Vcell throughout this book. A reaction diagram is constructed that connects all of the species.  Two are shown in Figure $7$ below. The computational results from the analyzes for yeast glycolysis were able to fit experimental data only if corrected by the addition of several branching reactions (shown in the figures above and Figure $8$) and in the dotted boxes in Figure 6 (Right) Trehalose is the non-reducing disaccharide Glc-(α 1,1)-Glc.  It is a reserve carbohydrate that also protects yeast against the effects of stress (desiccation, dehydration, temperature extremes) as well as lethal levels of ethanol.  These effects arise from its effects on protein stability which likely arise by its preferential exclusion from the hydration sphere of protein. A similar mechanism accounts for the stabilizing effect of glycerol on protein stability, as described in Chapter 4.9. c. A list of all reactions as shown in Figure $10$ for glycolysis and branching reactions (taken from COPASI). d. Parameters (concentrations, rate, enzyme kinetic, and equilibria constants) of all species. Figure $11$ below shows the initial concentrations for the glycolysis model. e. Equations that can be used to compute the change in concentrations of all species with time. These are usually ordinary differential equations (ODE) as described in Chapter 6B - Kinetics of Simple and Enzyme-Catalyzed Reactions, Sections B1: Single Step Reactions.) ODEs are easy to write but require a computer to solve as the number of interacting species increases. For a quick review, consider the relatively simple reaction scheme shown in Figure $12$ below. The set of ODEs for each species is shown below that. This set of ODEs for each species can be written. \begin{aligned} \frac{d[A]}{d t} &=-k_1[A][B]+k_2[C] \ \frac{d[B]}{d t} &=-k_1[A][B]+k_2[C] \ \frac{d[C]}{d t} &=+k_1[A][B]-k_2[C]-k_3[C]=+k_1[A][B]-\left(k_2+k_3\right)[C] \ \frac{d[D]}{d t} &=+k_3[C] \end{aligned} If a reaction removes species X, the right side of the ODE for the disappearance of that species has a – sign for that term. Likewise, if a reaction increases species X, the right-hand side has a + sign. The above examples show unimolecular (C to D, C to A + B) and bimolecular (A+B to C) reactions. Now let's look at the equations just for the change in the concentration of glucose-6-phosphate. One reaction forms it and 3 remove it, as shown in Figure $13$ below. Figure $13$: The reactions that change [G6P] in the Teusink yeast glycolysis model An ODE can be written for the change in concentration of glucose-6-phosphate. \left.\mathrm{d}[\mathrm{G} 6 P] / \mathrm{dt}=v_{\mathrm{HK}}-v_{\mathrm{PGI}}-2 v_{\text {trehalose }}-v_{\text {glycogen }}\right) where HK is hexokinase and PGI is phosphoglucoisomerase. Let's look at the terms more closely and how the terms are written in Vcell. Formation of G6P Only one reaction goes towards G6P.  Its the reaction catalyzed by hexokinase (G + ATP →  G6P + ADP).  In the Vcell reaction diagram, ATP is represented by P.  Flux J = \frac{V m G L K \cdot\left(G L C i \cdot A T P-\frac{G 6 P \cdot A D P}{K e q G L K}\right)}{K m G L K G L C i \cdot K m G L K A T P \cdot\left(1.0+\frac{G L C i}{K m G L K G L C i}+\frac{G 6 P}{K m G L K G 6 P}\right) \cdot\left(1.0+\frac{A T P}{K m G L K A T P}+\frac{A D P}{K m G L K A D P}\right)} Removal of G6P 3 reactions go away from G6P. i.  G6P → F6P.    Flux J = \frac{V m P G I_{-} 2 \cdot\left(G 6 P-\frac{F 6 P}{K e q P G I_{-} 2}\right)}{K m P G I G 6 P_{-} 2 \cdot\left(1.0+\frac{G 6 P}{K m P G I G 6 P \_2}+\frac{F 6 P}{K m P G I F 6 P_{-}2}\right)} ii.  G6P → Glycogen Flux J = KGLYCOGEN_3 iii.  G6P → Trehalose Flux J = KTREHALOSE Now let's run a simulation of the model of yeast glycolysis (Teusink,B et al.: Eur J Biochem 2000 Sep;267(17):5313-29).  The actual model is available from Biomodels (BIOMD0000000064). Yeast Glycolysis Yeast Glycolysis Teusink et al., 2000.  BIOMD0000000064 For Figure $14$ below, the model above model was run and the CSV file was used to plot the concentration vs time plots for each species.  Two of the species, EtOH (50 mM) and GLCo - glucose outside (50 mM), have fixed concentrations much larger than the rest, so they are omitted from the graph for clarity. When you run the simulation above, do the same when you download the csv file and plot the progress curves.  Also, note that when you run the simulation, the y-axis values are not shown correctly.  Rescale them in the spreadsheet based on the correct values of EtOH and GLC0, which were held constant (clamped) throughout the time course at 50 mM. Figure $14$: Time course graphs of all glycolytic species vs time How close does simulation replicate fluxes found experimentally in vivo?  As mentioned above, without the additions of the branch reaction, steady state was not achieved in the simulation. Fluxes in the model were within a factor of two for only about half of the enzymes so the model needs improvement.  Other discrepancies could be accounted for generically by differences in kinetics in vivo compared to in vitro measurements.  Additional products (glycogen, trehalose, glycerol and succinate) were derived from glucose, which mainly entered glycolysis and was converted to ethanol, showing the complexity in modeling even a relatively "simple" pathway as glycolysis. Metabolic Control Analysis and Simple Enzyme Inhibition Biochemists model complex enzyme-catalyzed reactions in the presence and absence of modifiers (either activators or inhibitors) to develop mechanisms for the reactions. The following kinetic parameters are experimentally determined by fitting initial velocity (vo) vs substrate concentration ([S]) through nonlinear fitting algorithms. • Km – the Michaelis Constant, the concentration of substrate at half-maximal velocity; • Vm – the velocity at saturating substrate concentration; • kcat - the turnover number for conversion of bound reactant to product; • Kix – inhibition dissociation constants. An example of competitive inhibition, shown in Figure $15$, illustrates a common type of analysis for such reactions. One modern pictorial depiction of a simple irreversible, enzyme-catalyzed reaction of substrate S going to product P with inhibition by the product and by an added inhibitor is shown in Figure $16$. The square, a "node" in the reaction diagram, represents the enzyme. Consider the simple enzyme-catalyzed reaction for a reversible conversion of substrate S to product P that has 3 reversible steps., as shown in Figure $17$ If the forward (f) and reverse (r) chemical reaction steps were irreversible and written separately, simple Michaelis-Menten equations could be written for each. \begin{aligned} &v_f=\frac{V_f S}{K_{M S}+S}=\frac{\frac{V_f S}{K_{M S}}}{1+\frac{S}{K_{M S}}} \ &v_r=\frac{V_r P}{K_{M P}+P}=\frac{\frac{V_r P}{K_{M P}}}{1+\frac{P}{K_{M P}}} \end{aligned} For the actual reversible reactions, the net forward rate cannot be found by simple subtraction of the two equations above as the differential equations describing the simple forward and reverse rates don’t account for the reverse steps v \neq\left[\frac{\frac{V_f S}{K_{M S}}}{1+\frac{S}{K_{M S}}}-\frac{\frac{V_r P}{K_{M P}}}{1+\frac{P}{K_{M P}}}\right] A simple derivation (assuming rapid equilibrium for both forward and reverse steps) can be made for the net forward reaction. Again consider the following enzyme-catalyzed reaction (Figure 17 above): A derivation of the rapid equilibrium, fully reversible Michaelis-Menten equation conversion of substrate S to product P Here it is! Derivation You may remember that for the isolated E + S ↔ ES and for the E + P ↔ EP reactions, the simple dissociation constants, KS and KP are given by \begin{aligned} &K_S=\frac{E_{e q} S_{e q}}{E S_{e q}}=\frac{k_{-1}}{k_1} \text { or } E S=\frac{[E][S]}{K_S} \ &K_P=\frac{E_{e q} P_{e q}}{E P_{e q}}=\frac{k_3}{k_{-3}} \text { or } E P=\frac{[E][P]}{K_P} \end{aligned} The rapid equilibrium assumption states that the rate of dissociation of ES and EP, which are both physical steps, is much faster than the rate of the chemical conversion steps for each complex. Hence $k_{-1} \gg k_2$ and $k_3 \gg k_{-2}$, so the relative amounts of ES and EP can be determined from the dissociation constants as shown above. Mass conservation of enzymes gives E_0=E+E S+E P=E+\frac{[E][S]}{K_S}+\frac{[E][P]}{K_p}=E\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right] From this, we can get the fractional amount of both ES and EP \begin{aligned} &\frac{E S}{E_0}=\frac{\frac{[E][S]}{K_S}}{E\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right]} \text { or } E S=\frac{E_0 \frac{[S]}{K_S}}{\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right]} \ &\frac{E P}{E_0}=\frac{[E][P]}{E\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_P}\right]} \text { or } E P=\frac{E_0 \frac{[P]}{K_P}}{\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_P}\right]} \end{aligned} Now we can derive the rate equation for the net forward reaction for the rapid equilibrium case: v=k_2[E S]-k_{-2}[E P]=k_2 \frac{E_0 \frac{[S]}{K_S}}{\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right]}-k_{-2} \frac{E_0 \frac{[P]}{K_p}}{\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right]} Knowing that k2E0 and k-2E0 represent the maximal velocities, Vf and Vr, respectively, the equation becomes: v=k_2[E S]-k_{-2}[E P]=\frac{V_f \frac{[S]}{K_S}}{\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right]}-\frac{V_r \frac{[P]}{K_p}}{\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right]}=\frac{V_f \frac{[S]}{K_S}-V_r \frac{[P]}{K_p}}{\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right]} Here is the derived equation. v=k_2[E S]-k_{-2}[E P]=\frac{V_f \frac{[S]}{K_S}}{\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right]}-\frac{V_r \frac{[P]}{K_p}}{\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right]}=\frac{V_f \frac{[S]}{K_S}-V_r \frac{[P]}{K_p}}{\left[1+\frac{[S]}{K_S}+\frac{[P]}{K_p}\right]} An equation of a similar form can be derived from the steady state assumption. Hence the net forward rate can't be determined by simple substration of the backward rate from the forward rate as shown in Equation 10 above. Common Forms of Kinetic Equations Programs like COPASI and VCell have many built-in equations for the velocities of many enzyme-catalyzed reactions that have similar forms. Two are shown below: Reversible Michaelis-Menten: \frac{\frac{V_f[\text { substrate }]}{K_{M s}}-\frac{V_r[\text { product }]}{K_{M p}}}{1+\frac{[\text { substrate }]}{K_{M s}}+\frac{[\text { product }]}{K_{M p}}} Competitive Inhibition Reversible: \frac{\frac{V_f[\text { substrate }]}{K_{M s}}-\frac{V_r[\text { product }]}{K_{M p}}}{1+\frac{[\text { substrate }]}{K_{M s}}+\frac{[\text { product }]}{K_{M p}}+\frac{[\text { inhibitor }]}{K_I}} What is most important for readers to understand is not detailed derivations or how to solve the differential equations on their own. However, you should be able to: • write the differential equation of a given reaction; • recognize the common equations used for nonenzyme-catalyzed reactions (mass action) and enzyme-catalyze ones; • change parameters in programs that use numerical methods to solve systems of linked differential equations for a system and see how the time course graphs are affected.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/14%3A_Principles_of_Metabolic_Regulation/14.02%3A_Basic_Principles_of_Metabolic_Control_Analysis_%28MCA%29.txt

Search Fundamentals of Biochemistry Introduction Metabolic control analysis (MCA) is one method used to address the complexity of dynamic changes of species in a complex metabolic system. As such, an understanding of MCA would apply to complex signal transduction pathways as well as to other emerging areas in systems biology. In MCA, external inputs (source) and outputs (exits), and pools and reservoirs exist which are connected to the internal metabolic enzymes, reactants, and products of the pathway connecting the two external reservoirs. In studying complex metabolic systems, what is interesting to know is not just the $K_m$ or $V_m$ of particular enzymes (such as those far from equilibrium or catalyzing “rate limiting” steps based on in vitro studies), but rather the control parameters, known as control coefficients, for the system. These coefficients are properties of the whole pathway as an emerging system with properties different from those of the individual steps. For another example of an emergent property, consider consciousness. It must emerge from the incredible assembly of around 90 billion neurons in the brain forming around 1015 synapses.  It cannot simply be predicted from the properties of isolated neurons. Three especially relevant parameters are used in metabolic control analyses, the Flux Control Coefficient, the Concentration Control Coefficient, and the Elasticity Coefficient To understand how a pathway is regulated and how scientists modify it to meet their demands, one needs to understand these coefficients.  In particular, we would like to know which: • parameters have the greatest effect (increase or decrease) on a preferred outcome; • properties of the system make it most resilient or fragile; • parameters should be measured most accurately The Flux Control Coefficient The Flux Control Coefficient ($C_{Ei}^{J}$) gives the relative fractional change in pathway flux $J$ ($dJ/J$) (a system variable) with fractional change in concentration or activity of an enzyme $Ei$ ($du_i/u_i$). The change in the concentration of an enzyme can arise from increased synthesis or degradation of the enzyme, a change in a modifier (an inhibitor or activator), a covalent modification, the presence of inhibitor RNA, mutations, etc. Again, the word flux $J$ is used to describe the rate of the system whereas rate or velocity $v$ is used to describe an individual enzyme in the system. Think of $u$ as “units” of activity”. The Flux Control Coefficient ($C_{Ei}^{J}$) can also be written as $C_{Ei}^{J}=\dfrac{d J}{d u} \dfrac{u}{J} \label{15}$ The $dJ/du$ in Equation \ref{15} suggests that the $C_{Ei}^{J}$ is equal to the slope of a plot of $J$ vs u multiplied by $u/J$ at that point. Alternatively, it is the actual slope of an ln-ln plot of $\ln J$ vs $\ln u$. A more formal definition would be obtained using partial derivatives (a normal derivative like dy/dx with everyother variable held constant - in this case all other enzyme concentrations or activities other than the one under study : $C_{E i}^{J}=\dfrac{\dfrac{\partial J}{J}}{\dfrac{\partial u}{u}}=\dfrac{\partial \ln J}{\partial \ln u_{i}} \label{16}$ $C_{Ei}^{J}$ clearly shows how the system flux changes with changes in the activity of a single enzyme. The variable ui is a property of an enzyme (a local property) but J and concentrations (for the concentration coefficient – see below) are systems variables. Every enzyme in a pathway hence has a flux control coefficient. How are they related? It makes sense that the sum of all of the individual $C_{Ei}^{J}=1$ as the fractional change in flux, $∂J/J$, is equal in magnitude to the fractional change in all enzyme activities. This relationship, shown below, is called the Summation Theorem. $\sum_{i=1}^{n} C_{E i}^{J}=1 \label{17}$ This equation would predict that if there were one magical enzyme X that was truly rate limiting and as such completely controlled that rate, then CJEx = 1 for that enzyme and 0 for all the rest of the enzymes. Yet this is counterintuitive since if all the enzymes are linked their responses should also be linked. Hence the control of flux is shared by all enzymes. Another way to think about this is to consider a linear set of connected reactions. For these enzymes, $C_{Ei}^{J}$ would vary between 0 and 1. If one enzyme was completely rate limiting, its $C_{Ei}^{J}$ would be 1 indicating that the fractional change in flux, $∂J/J$, is equal to the fractional change in enzyme concentration (or activity) $u$, $∂u/u$, for enzyme X. This implies that all change in flux is accounted for by the change in the concentration of enzyme X. This is a bit difficult to believe so it would make sense to think of the control of a pathway as being distributed over all the enzymes in the pathway. Plots of flux $J$ vs enzyme activity u are similar to rectangular hyperbolas. Hence the flux coefficient changes with enzyme activity and with flux so at different fluxes, different enzyme flux coefficients would change as well. “Control” gets redistributed in the pathway. Textbooks often describe that ideally the first step of a linear pathway would be the committed step and be rate limiting. But think of this. If the end product of a pathway feeds back and inhibits the first enzyme in the pathway, then the last enzyme influences the first even though the last is not considered “rate limiting”. Now if another enzyme removes the last product and hence first enzyme's feedback inhibitor of the pathway, that enzyme determines flux as well. This again shows that flux control is distributed throughout the system. The classical “rate limiting” enzyme might still have the largest $C_{Ei}^{J}$ of the pathways, but if the system is in a steady state, all enzymes have the same rate. Hence the term “rate-limiting” is a bit useless in a pathway. Analysis of flux control coefficients gives a truly quantitative analysis of the system pathway. Phosphofructokinase is said to be a “key” enzyme in glycolysis as it is one of three enzymes in the pathway that is characterized by a large negative free energy difference.   Figure $1$: Figure $1$: ΔG0 and ΔG values for enzymes in glycolysis It is also highly regulated. However, Heinisch et al (Molecular and General Genetics, 202, 75-82) found that increasing its concentration 3.5 fold in fermenting yeast did not affect the flux of ethanol production. Here are some results of "sensitivity analyses" of yeast glycolysis showing the effect of small changes (up to 5%) on the flux control coefficients of glycolytic enzymes. The changes in the enzymes for these analyses are 1%. The numbers are scaled values that represent a relative change. A value of + 0.5 means that if the parameter is increased by 10%, the target value will increase by 5%. A column shows the rate of a reaction that is changed and the row indicates the flux of the reaction that has been affected. Green represents positive values and red negative with the intensity of the color correlating with the extent of the change. The table is split with column 1 of the first table applying to tables 2 and 3 as well. Table, part 2, continued: table, part 3, continued Some things to note: • almost everything has changed in each column, which shows that a change in one enzyme affects all enzymes; • 2 adjacent rows often have identical numbers since they describe two connected reactions (without branches) so their steady state fluxes will be the same; • the sum of the CJEi values across the same row in both tables is 1 (summation theorem); • perturbations of the "big 3" enzymes for steps 1,3, and 10 in glycolysis (hexokinase, phosphofructokinase, and pyruvate kinase), all of which proceed with a significant -ΔG0 and -ΔG, have minor effects on the flux. It should be clear from these results that you can't predict the enzymes that control the flux without this kind of mathematical analysis. Don't rely on just your "biochemical intuition" Flux Control in Aerobic Glycolysis Let's look at another pathway for which we can run a full simulation using Vcell - aerobic glycolysis.  We have mostly encountered glycolysis as a central anaerobic pathway in almost all organisms.  In some cases, it can also run aerobically.  One prime example is in cancer cells in which there is a need for energy in the form of ATP but also for anabolic building blocks so cells can proliferate - a hallmark of cancer cells.  Aerobic glycolysis was noted by Warbug and this effect now has his name. Aerobic glycolysis has been modeled in yeast cells.  The pathway is shown in Figure $2$ below. Figure $2$:  Schematic of the glycolysis model with chemical reactions and allosteric points of regulation described. Alexander A Shestov et al.  (2014) Quantitative determinants of aerobic glycolysis identify flux through the enzyme GAPDH as a limiting step. eLife 3:e03342.  https://doi.org/10.7554/eLife.03342.  Creative Commons Attribution License Abbreviations: GLC—glucose, G6P—glucose-6-phosphate, F6P—fructose-6-phosphate, FBP—fructose-1,6,-bisphosphate, F26BP—fructose-2,6,-bisphosphate, GAP—glcyceraldehyde-3-phosphate, DHAP—dihydroxyacetone phosphate, BPG—1,3 bisphosphoglycerate, 3PG—3-phosphoglycerate, 2PG—2-phosphoglycerate, PEP—phosphoenolpyruvate, PYR—pyruvate, SER—Serine, GLY—glycine, Lac—lactate, MAL—malate, ASP—aspartate, Pi—inorganic phosphate, CI—creatine, PCI—phosphophocreatine, GTR—glucose transporter, HK—hexokinase, PGI—phosphoglucoisomerase, PFK—phosphofructokinase, ALD—aldolase, TPI—triosephosphoisomerase, GAPDH—glyceraldehyde-phosphate dehydrogenase, PGK—phosphoglycerate kinase, PGM—phosphoglycerate mutase, ENO—enolase, PK—pyruvate kinase, LDH—lactate dehydrogenase, MCT—monocarboxylate transporter, PDH—pyruvate dehydrogenase, CK—creatine kinase. The Warburg Effect W can be quantitatively described by the ratio of the flux of pyruvate to lactate (JLac) compared to the flux of pyruvate entry into the mitochondria and subsequent consumption of O2  by mitochondrial oxidative phosphorylation (Jox), so W = JLac/Jox). Usually in normal conditions for healthy cells, W <0.10, which means that less than 10% of glucose is converted (or diverted) to lactate.  On average for different tissues, W < 0.3.  Hence some lactate is always being made.  High values of W (> 1) mean that most glucose is diverted to lactate synthesis. The W values is determined by lots of factors that determine the energy state of cells, including rates of glucose uptake, ATP hydrolysis, and biosynthesis, as well as the balance of cytoplasmic and mitochondrial NAD+/NADH, hence the redox state of the cell. Key to the balance leading to aerobic glycolysis is the need to reoxidize cytosolic NADH back to NAD+ under aerobic conditions so glycolysis can continue under conditions when there is a high growth rate. How does a cell know which path to take, pyruvate to lactate (anaerobic), pyruvate to CO2 with ATP synthesis (aerobic), or some of both?  In terms of metabolic control analysis, what controls the flux of pyruvate through these paths?  This is where computational modeling informed by experimental data helps. We can determine the flux control coefficient ($C_{Ei}^{J}$),  more easily written as FCC, for each enzyme in glycolysis to help us understand what controls the flux.  Figure $3$ metaphorically describes these different paths. In a way, the flux of pyruvate through these 3 choices, anaerobic, aerobic, and both (as reflected by the Warburg effect W) is then perhaps the best example to use to discuss flux control since the readers are likely very familiar with the concept of anaerobic and aerobic metabolism and the "switch" between them when running short sprints vs marathons. Figure $3$: Fluxs of pyruvate through multiple paths Now let's run the Vcell model for aerobic glycolysis in yeast. MODEL AerobicGlycolysis Quantitative determinants of aerobic glycolysis identify flux through the enzyme GAPDH as a limiting step. .Shestov AA, Liu X, Ser Z, Cluntun AA, Hung YP, Huang L, Kim D, Le A, Yellen G, Albeck JG, Locasale JW.  eLife , 7/ 2014 , Volume 3 , PubMed ID: 25009227.  Biomodel MODEL1504010000 Select Load [model name] below Select Start to begin the simulation. Interactive Element Select Plot to change Y axis min/max, then Reset and Play  |  Select Slider to change which constants are displayed |  Select About  for software information. Move the sliders to change the constants and see changes in the displayed graph in real-time. Time course model made using Virtual Cell (Vcell), The Center for Cell Analysis & Modeling, at UConn Health.  Funded by NIH/NIGMS (R24 GM137787); Web simulation software (miniSidewinder) from Bartholomew Jardine and Herbert M. Sauro, University of Washington.  Funded by NIH/NIGMS (RO1-GM123032-04) Where did all the parameters in the model come from?  They are derived experimentally and the goal of modeling is to produce a computational model that is consistent with the experimental findings which are fed into the model. The result of the model suggests that the Warburg effect is determined by the growth rate of the cells as well as the activity of mitochondria. The model does not support the idea that aerobic glycolysis results as a balance between energy needs and new biosynthesis as cells proliferate. Rather, the cells that proliferate the most have the most active mitochondria and hence lower levels of lactate production.   Redox balance appears to be key in pushing cells toward lactate synthesis and aerobic glycolysis. A key step is catalyzed by glyceraldehyde-3-phosphate dehydrogenase (GAPDH), the glycolytic enzyme that separates the top and bottom halves of glycolysis. Experimental data shows that in fact, flux through this enzyme is rate-limiting with the levels of F1,6-BP also being very important. Some key steps that were thought to be rate-limited in fact had negative fluxes through them. Negative flux control coefficients were found and confirmed for several steps thought to be rate-limiting in glycolysis. The flux control coefficients, given by the formula C_{E i}^J=\frac{\frac{\partial J}{J}}{\frac{\partial u}{u}}=\frac{\partial \ln J}{\partial \ln u_i} were calculated for each enzyme in the glycolytic pathway.  The authors of the study used this equation for the Flux Control Coefficient: F C C=\frac{d \ln \left[J_{L A C T}\right]}{d \ln \left[E_i\right]} The fluxes were measured for lactate production by using 13C-labeled lactate to get the absolute concentration of lactate, and then measuring the fluxes by measuring the changes in 13C-lactate production with time. Using these data and the selective inhibition by small drugs of each step in glycolysis to effectively alter the "concentration/activity of the enzyme", the FCC of each enzyme could be calculated.  The results are shown in Figure $4$ below. Figure $4$: (left) Box plots of flux control coefficient (FCC) for lactate production for each enzymatic step in glycolysis (FCC = dlnJlac/dln Ei) where Jlac is the rate of pyruvate conversion to lactate, and Ei is the ith enzyme in glycolysis for each step of glycolysis. (right) Box plots of flux control coefficient (FCC) for lactate production for Oxygen consumption (OxPhos) and ATP consumption (ATP). Shestov et al., ibid The left-hand side of Figure 4 shows that two of the key enzymes that are thermodynamically favored in glycolysis, PFK (phosphofructokinase) and HK (hexokinase), both driven by ATP hydrolysis, have essentially negative flux coefficients, so they are NOT key in driving flux toward lactate.  Only one enzyme, GAPDH, is characterized by a positive flux coefficient. (Remember we are talking about flux coefficients, not the Warburg effect factor W.) Also important, as mentioned above, is the concentration of F1,6BP (FBP in the figure below).  The model and data show that high levels of FBP and associated higher concentrations of the reactants in the first half of glycolysis (denoted by ↑↑ and large font size), and a corresponding depletion of substrates in the bottom half (denoted by ↓↓) actually cause a bottleneck in glycolysis and inhibit flux through GADPH.  Hence GADPH determines the flux through glycolysis. This "bottleneck" is associated with the redox and energy states of the cell.   This outcomes are illustrated in Figure $1$ below. Figure $5$: Figure $5$:  A unified model of aerobic glycolysis. Shestov et al., ibid When there is a balance of metabolites in the top and bottom halves of glycolysis (right hand size), including both high levels in both top and bottom (denoted by ↑↑ and font size) or low levels in both (denoted by ↓↓ and font size), no bottleneck at GAPDH exists and flux through glycolysis is determined by the usual "suspects", hexokinase and phosphofructokinase. The results described above are those predicted by computational modeling.  Do experimental data support the model?  The only way to know is to test the model.  This was done by using drugs to inhibit each of the enzymes in glycolysis.  This offers an effective way to change the "effective" concentration or activity in the enzyme.  From this data, flux changes due to changes at each step could be determined. These data supported the computation results - GADPH regulates aerobic metabolism in normal as well as.  When the top half substrates were elevated, the blockade of flux was determined by GADPH.  In Chapter 13, we studied all of the glycolytic enzymes and the myriad of small molecules that control the activity of the 3 major enzymes that have the biggest thermodynamic push, HK, PFK and PK.  GADPDH appears to be equally important.  It is known to be regulated by many mechanisms, including post-translational modifications, such as nitrosylation, and modification of its active site cysteine with ROS.  Perhaps this helps regulate the balance between anaerobic and aerobic glycolysis.  Also, there are high levels of expression of GADPH in cells engaged in aerobic glycolysis.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/14%3A_Principles_of_Metabolic_Regulation/14.03%3A_The_Flux_Control_Coefficient.txt

Search Fundamentals of Biochemistry Concentration Control Coefficients The Concentration control coefficient ($C_{E_{i}}^{S}$), a global property of the system, gives the relative fractional change in metabolite concentration $S_j (dS_j/S_j)$, where $S_j$ is the concentration of any metabolite in the system and as such is a system variable, with fractional change in concentration or activity of enzyme $E_i (du_i/u_i)$. Similar equations as section 15.3 apply. C_{E_i}^S=\frac{\frac{\partial S_i}{S_i}}{\frac{\partial u_i}{u_i}}=\frac{\partial \ln S_i}{\partial \ln u_i}=\frac{\partial S_i}{\partial u_i} \frac{u_i}{S_i} It can be shown that the sum of all of the individual $C^S_{E_i} = 0$ (another summation theorem), is not 1 as in the case of flux control coefficients. This again would make sense in the steady state. Flux coefficients usually vary from 0 to 1, but concentration coefficients can vary from negative to positive and small to large. \sum_{i=1}^n C_{E i}^S=0 The concentration control coefficients can have large values as seen in a simple example. For a given enzyme, at low [S], for example, when [S] << Km, v=\frac{V_m S}{K_M+S}=\frac{V_m S}{K_M}=\frac{k_{c a t} E_{t o t} S}{K_M} \text { when } S \ll K m If the enzyme had only 0.1x of its normal activity (due to a mutation for example), then to maintain constant flux, the [S] would have to increase 10-fold. The tables below show the CSEi values for incremental changes in the substrate (1%). table, part 2: table, part 3 Elasticity Coefficient The Elasticity Coefficient, in contrast to the flux and concentration control coefficients, which are properties of the system, is a local property and can be measured using isolated enzymes and substrates. It makes sense that some kinetic property of the isolated enzyme would affect its propensity to affect system flux. The elasticity coefficient gives a measure of how much a substrate $S$ (or other substance) can change the reaction rate ($v$) of an isolated enzyme. (Note we use $v$ and not flux $J$, which describes a system property.) Hence \varepsilon_S^v=\frac{\frac{\partial v}{v}}{\frac{\partial S}{S}}=\frac{\partial \ln v}{\partial \ln S}=\frac{\partial v}{\partial S} \frac{S}{v} Price Elasticity The term elasticity is also used in economics and is especially useful in times when inflation is high. If the price of your favorite product, such as a Starbucks Latte coffee goes up, consumers might either buy them less frequently or buy another cheaper latte from a competitor. If a small rise for a Starbucks latte leads to a large drop in demand, the Starbucks latte is characterized by a high elasticity. If however, people don't change their latte buying behavior when there is a big price rise on the latte, the product is said to be inelastic. If you are the CEO of a company, it is good to know the elasticity of your products to maximize your profits. Hence the elasticity coefficient can be determined using basic enzyme kinetics of the isolated enzyme. Note that the coefficient at each $S$ concentration is the slope of the v vs S curve multiplied by the $S/v$ at that tangent point. The elasticity coefficient must be evaluated at the same concentration of enzyme and substrate as found in vivo in the steady state. Velocity ($v$), not flux, is used. In the above case, $S$ is the substrate, but it could be a product or modifier. There is a different elasticity coefficient for each parameter. There is no summation theory for elasticities. Values can be positive for species that increase the velocity or negative for those that decrease it. Hence there can be multiple elasticities. The tables below show the elasticity coefficients (relative or scaled) for yeast glycolysis determined using COPASI. table, part 2 Things to note: • the columns show substrates not enzymes; • green cells (with positive elasticities) are generally substrates for their target enzymes (for example, glucose-6-phosphate for phosphoglucomutase); Other "generic" sensitivities can be determined as well. For example, incremental changes in $K_m$, $V_m$, or $K_{ix}$ for specific enzymes could affect fluxes in a pathway. A link between system control coefficients and local coefficients: You would think that there should be some relationship between a system variable such as the flux control coefficient and a local variable such as the elasticity coefficient. There is and it is expressed by the Connectivity Theorem (below). If a substrate $S$ is acted upon by many different enzymes ($i …… n$), which is very likely, especially for branch points in metabolic pathways, then it can be shown that \sum_{i=1}^n C_{v i}^J e_s^{v i}=0

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/14%3A_Principles_of_Metabolic_Regulation/14.04%3A_Concentration_Control_and_Elasticity_Coefficients.txt

Search Fundamentals of Biochemistry Introduction We have studied binding interactions in Chapter 5, kinetics in Chapter 6, and principles of metabolic control in this chapter.  We've learned the following: Binding Reactions • for simple binding of a ligand to a macromolecule, graphs of fractional saturation of the macromolecule vs free ligand concentration are hyperbolic and demonstrate saturation binding.  In the initial part of the binding curve, when [L] << KD, the fractional saturation shows a linear dependence on free ligand concentration.   Figure $1$ shows [ML] vs L, which is the same basic equation as a plot of Y vs L. Figure $1$ • for allosteric binding of a ligand to a multimeric protein, graphs of fractional saturation vs free ligand concentration are sigmoidal and also display saturation binding. In the first parts of the binding curve, the fractional saturation is much more sensitive to ligand concentration than in simple binding of a ligand to a macromolecule with one binding site.  Figure $2$ below shows graphs for the allosteric binding of a ligand to a macromolecule using the Hill Equation (instead of the MWC equation we used to model O2 binding to tetrameric hemoglobin). Figure $2$ In these two plots, the system (in this case a single macromolecule) displays different sensitivities to ligand concentration, allowing the system to have different responses to changes in physiological conditions. Binding and Chemical Reactions As with the case for binding interactions, we have seen hyperbolic and sigmoidal plots of initial velocity (v0) vs [substrate] for enzyme-catalyzed reactions. These also allow appropriate responses to a single substrate in a physiological setting. But what if you put the same macromolecule and ligand into a larger metabolic or signal transduction pathway in vivo? What kinds of responses would they make to a change in input?  As we have just seen in our discussion of the steady state, the ligand or substrate concentration might not change at all as flux continues through the pathway. One could imagine a lot of scenarios with different inputs and different optimal outputs.   For example, what if the input (a reactant or small signaling molecule) comes in pulses? Ultimately a system should return to its basal state since a prolonged response (such as cell proliferation) could be detrimental to the health of the organism. Let's look at some simple examples and see how different inputs lead to specific outputs.  We'll just construct some very simple reaction diagrams in Vcell and see how varying them leads to different outputs.  Here are two simple cases for isolated chemical species and reactions, analogs to the simple binding reactions described above. Linear Response: A Signal S and a Response R;  S → R If no enzyme is involved, the rate doubles as the signal (substrate) doubles since dR/dt = k[S] for the first-order reaction.  If S is the stimulus and R is the response,  a plot of R vs S is linear.  Hence the system responds linearly with increasing S. Here is the simple chemical equation \mathrm{S} \underset{\mathrm{k}_2}{\stackrel{\mathrm{k}_1}{\rightleftarrows}} \mathrm{R} As a concrete example, consider the synthesis and degradation of a protein, characterized by the following equation derived from mass action. \frac{d R}{d t}=k_0+k_1 S-k_2 R where S is the signal (ex. concentration of mRNA) and R is the response (ex concentration of the transcribed protein).  A constant k0 has been added to account for any basal rate of the reaction.  (This is a vastly oversimplified way to model a complex process like mRNA translation to a protein as it omits 100s of steps.) Here is the simplified derivation under steady state (SS) conditions typically found for enzymes embedded in a pathway. \begin{gathered} \frac{d R_{S S}}{d t}=k_0+k_1 S-k_2 R=0 \ R_{S S}=\frac{k_0+k_1 S}{k_2} \end{gathered} The equation is a linear function of S. Hyperbolic Response:  E+S ↔ ES → E + R In a simple enzyme-catalyzed reaction with a fixed concentration of enzyme, as S increases the initial velocity saturates.  Hence there is a limit on the response, so the response R is a hyperbolic function of S.  Increasing S ever more after saturation won't lead to more R (in a given amount of time). As a concrete example of this consider the phosphorylation/dephosphorylation of a protein R.  RP represents the phosphorylated and active form of the protein R with concentration [RP].  The reaction is simply written as R ↔ RP, where RP is the response.  Mass action shows that the total amount of R, RT = R + RP.   A simple mass action equation can be derived. Here is the chemical equation \mathrm{R}+\mathrm{S} \underset{\mathrm{k}_2}{\stackrel{\mathrm{k}_1}{\rightleftarrows}} \mathrm{R}_{\mathrm{P}} Here is the math equation, again for the steady state (SS), when dRP/dt = 0.  (We derived the same equation for the steady-state version of the Michaelis-Menten equation in Chapter 6. \frac{d R_P}{d t}=k_1 S\left(R_T-R_P\right)-k_2 R_P Derivation: Steady State Hyperbolic Response to a Stimulus S Click below to see the derivation Derivation \frac{d R_P}{d t}=k_1 R[S]-k_2 R_P then in the stead state: \begin{gathered} \frac{d R_P}{d t}=k_1 S\left(R_T-R_P\right)-k_2 R_P=0 \ k_2 R_{P, S S}=k_1 S\left(R_T\right)-k_1 S\left(R_{P, S S}\right) \ k_2 R_{P, S S}+k_1 S\left(R_{P, S S}\right)=k_1 S\left(R_T\right) \ R_{P, S S}\left(k_2+k_1 S\right)=k_1 S\left(R_T\right) \end{gathered} Finally, we get R_{P, S s}=\frac{k_1 S\left(R_T\right)}{\left(k_2+k_1 S\right)}=\frac{\left(R_T\right) S}{\left(\frac{k_2}{k_1}+S\right)} In the steady state, dRP/dt = 0, and the steady state equation can be written as: R_{P, s s}=\frac{k_1 S\left(R_T\right)}{\left(k_2+k_1 S\right)}=\frac{\left(R_T\right) S}{\left(\frac{k_2}{k_1}+S\right)} Sigmoidal Response Consider this simple reaction for a homotetramer in which each monomer can bind a substrate S:  nS + En  ↔ EnSn → En + nR:  If En is a multimeric allosteric enzyme, as S increases the initial velocity also saturates but the response R is a sigmoidal function of S (in analogy to the above example).  The equation is too complicated to derive there, but the result reproduces a sigmoidal curve for the steady state, much as the Hill equation does for cooperative binding. Adaptation and Homeostasis The above examples show that the response of proteins or enzymes to increasing levels of a stimulus like a ligand or a substrate can be linear, hyperbolic, or sigmoidal, with quite a varied set of outcomes.  However, in many biological conditions, an ever-increasing or increasing and plateauing response might be too much.  The cell needs a way to turn off the response and settle back to a basal state, even in the presence of constant or changing stimuli.  This allows the adaption of a system to a stimulus and the maintenance of homeostasis. Every system needs to be able to respond and return to a homeostatic basal level.  The maintenance of homeostasis is critical to life. Homeostasis - ASBMB The American Association for Biochemistry and Molecular Biology (ASBMB) describes both homeostasis and evolution as key underlying concepts for all biology.  Homeostasis shapes both form and function from the molecular to organismal levels. Homeostasis is needed to maintain biological balance.  The steady state at the molecular to organismal levels in metabolic and signaling pathways is a hallmark of homeostasis. Here are the learning goals for homeostasis designated by the ASBMB 1.  Biological need for homeostasis Biological homeostasis is the ability to maintain relative stability and function as changes occur in the internal or external environment. Organisms are viable under a relatively narrow set of conditions. As such, there is a need to tightly regulate the concentrations of metabolites and small molecules at the cellular level to ensure survival. To optimize resource use, and to maintain conditions, the organism may sacrifice efficiency for robustness. The breakdown of homeostatic regulation can contribute to the cause or progression of disease or lead to cell death. 2. Link steady-state processes and homeostasis A system that is in a steady state remains constant over time, but that constant state requires continual work. A system in a steady state has a higher level of energy than its surroundings. Biochemical systems maintain homeostasis via the regulation of gene expression, metabolic flux, and energy transformation but are never at equilibrium. 3. Quantifying homeostasis Multiple reactions with intricate networks of activators and inhibitors are involved in biological homeostasis. Modifications of such networks can lead to the activation of previously latent metabolic pathways or even to unpredicted interactions between components of these networks. These pathways and networks can be mathematically modeled and correlated with metabolomics data and kinetic and thermodynamic parameters of individual components to quantify the effects of changing conditions related to either normal or disease states. 4. Control mechanisms Homeostasis is maintained by a series of control mechanisms functioning at the organ, tissue, or cellular level. These control mechanisms include substrate supply, activation or inhibition of individual enzymes and receptors, synthesis and degradation of enzymes, and compartmentalization. The primary components responsible for the maintenance of homeostasis can be categorized as stimulus, receptor, control center, effector, and feedback mechanism. 5. Cellular and organismal homeostasis Homeostasis in an organism or colony of single-celled organisms is regulated by secreted proteins and small molecules often functioning as signals. Homeostasis in the cell is maintained by regulation and by the exchange of materials and energy with its surroundings. In the rest of the chapter section, we will describe chemically and mathematically simple circuits/motifs that are employed that allow perfect or near-perfect adaptation to a stimulus, a hallmark of homeostasis.  We will define adaptation as a complete or almost complete return to a basal state after the introduction of a stimulus.  In all the cases below we will consider not a single application of a stimulus but a pulse application (a repetitive step wave function).  The pulsed stimuli could be of constant magnitude or an increasing/decreasing pulse of a signal such as a substrate.  All responses must be transient to avoid uncontrolled responses such as proliferation (a hallmark of tumor cells) or cell death. Adaptation is commonly found in sensory systems like vision, hearing, pressure, taste, etc.  Think of eating your favorite cookie. The first bite is delicious but by the tenth bite, there is significant attenuation in the positive sensory response, which helps keep most from adding significant weight continually. Ma et al. conducted simulations on three component/nodes (proteins, enzyme) systems to see which might display the potential for perfect or near-perfect adaption.  The simple 3-component motifs or circuits were modeled using simple mass action kinetic equations, ordinary differential equations (which we learned to write in Chapter 6.2), or a combination of both.  The systems that displayed adaption had to conform to three criteria: 1. The stimulus had to initially induce a response of high magnitude 2. The system had to return to a basal or near basal state. 3. The return to a basal state had to be mostly parameter-independent.  That is, the return to the basal state must occur for many different combinations of parameters. The possible 3-component components (nodes) and the links among the nodes are shown in Figure $3$ below. Figure $3$:  Possible 3-component components (nodes) and the links among the nodes.  After Ma et al. Cell Theory,138, 760-773 (2009) DOI:https://doi.org/10.1016/j.cell.2009.06.013. Out of over 16,000 models, several hundred were found that met the criteria. Most were variations of simple motifs that we will show below. The most common motifs were the negative feedback loop and the incoherent feedforward system Much of the discussions, models, and equations used below are from two articles: • John J Tyson, Katherine C Chen, Bela Novak, Sniffers, buzzers, toggles and blinkers: dynamics of regulatory and signaling pathways in the cell, Current Opinion in Cell Biology, Volume 15, Issue 2, 2003, Pages 221-231, https://doi.org/10.1016/S0955-0674(03)00017-6. • James E. Ferrell, Perfect and Near-Perfect Adaptation in Cell Signaling, Cell Systems, Volume 2, Issue 2, 2016, Pages 62-67, https://doi.org/10.1016/j.cels.2016.02.006. By adding a third component to form a mini pathway, we can now change the response R to a stimulus S from linear, or hyperbolic/sigmoidal in the steady state, to one that exhibits perfect or near-perfect adaptation. Again we see this kind of response in signaling pathways in sensation and also in responses like chemotaxis, in which a cell moves toward a stimulus (a chemoattractant molecule). Simple 3-node motif/circuit for perfect adaptation Figure $4$ below shows our first example of a 3-component system that displays perfect or near-perfect adaption.  The right-hand side shows a Vcell reaction diagram. In this example, a stimulus S (could be a reactant, neurotransmitter, mRNA, etc) leads to the synthesis of X and also of R, a response molecule.  Both X and R get degraded.  The yellow squares represent the nodes through which the flux of S to X and R proceeds.  Each node has an equation for the flux, J, through the node.  The left part of Figure 4 shows the periodic pulse of stimuli S that increases the concentration of S from an initial value of S0 = 1 uM to S + 0.2 uM for each step. Note that the flux equations for J are very simple and are based on mass action, and are not derived through Michaelis-Menten kinetic equations. Figure $4$:  Simple 3-component system that displays perfect or near-perfect adaption. Note that S, the stimulus (or substrate for example) is a square wave step function varying from 0 to 1 over the time interval shown in the graph.  The dotted blue line simply shows when the pulse is delivered.  The initial S concentration is 1 uM and increases by 0.2 for each step (as shown in the gray line).  Hence S increases in a stepwise fashion. Figure $5$ below is a time course graph that shows the stepwise (=0.2 uM) increase in S from 1 uM and the concentration of R (the response) over 20 seconds.  Even though S continues to increase in a stepwise fashion, R rises substantially only from the initial input of S (1 uM) and subsequent increases in S with each increment of S are damped out! Figure $5$: Time course for a 3-Component Perfect Response system.  Model by ModeBrick from VCell: CM-PM12648679_MB4:Perfect_Adaptation; Biomodel 188456707 Running Adaptive Vell models in this book The present version of Vcell release (as of 4/28/23) does not yet allow the export of a file compatible with the software used to run simulations with this book.  The Vcell model includes an "event" which allows for the production of stepwise changes in stimuli.  A future release will allow users to run the simulations within this book (as is the case for the other Vcell simulations throughout the book). Negative Feedback Loop The negative feedback loop is one of the simplest circuits/motifs to generate perfect or near/perfect adaptation.  It has only two nodes (yellow dots) and two proteins. An example is bacterial chemotaxis. Figure $6$ below shows a Vcell reaction diagram (left), and another representation (middle) and the time course graphs for all species. This model works especially well with certain parameters assigned. Figure $6$ Figure $6$:  Near-Perfect Adaptation from Negative Feedback. Adapted from Ferrell (ibid) The gray line in the graph is the stimulus S (substrate).  The blue line is the response, designated in this model as A.  B acts as an inhibitor (note the dotted line to the input node in the left diagram and the blunt-ended red bar in the middle diagram.  Note that the stimulus goes from 0.2 uM (initial concentration) at t=0 to 1 uM (a 5-fold increase) at 40 seconds, but the response A increases at most from 0.4 (initial condition) to 0.5 (a 1.25-fold increase). If we say the [A] is the output, then the differential equation for dA/dt is given by \frac{d A}{d t}=k_1 \operatorname{S} \cdot(1-A)-k_2 A \cdot B dB/dt is given by \frac{d B}{d t}=k_3 A \frac{1-B}{K_3+1-B}-k_4 \frac{B}{K_4+B} The constants for the graph (right) produced by the Vcell model are: • k1 = k2 = 200 • k3 = 10; k4 = 4 • K3 = K4 = 0.01 Incoherent Feedforward systems In this circuit/motif, the stimulus S increases the concentration of A (the output) but also forms a negative modulator, B, which with a bit of a time lag decreases the concentration of A through inhibition.  There is no feedback inhibition from A in this simple system. If you're reading carefully, you'll see that the reaction scheme and inhibition are the same as the first circuit/motif we introduced.  Here we simplify the diagram and give it an official name.  The word incoherent in the name makes sense since the stimulus S is converted both to the output A, and to the inhibitor B, which on the surface seems like a crazy thing to do. Figure $7$ below shows the Vcell reaction diagram (left) and more classical reaction diagram (middle) and progress curves showing S, the stimulus, A the output or response, and B, the inhibitor.  The dashed line in the left diagram from B to the reaction node for the S → A reaction shows that B affects the rate of that reaction. The equations used account for the inhibitory effect of B. Figure $7$:  Near-Perfect Adaptation from an Incoherent Feedforward System. Adapted from Ferrell (ibid) Note that the response A goes up or down a bit with each new step in concentration of S but to a very minimal degree. The system is certainly almost perfectly adapted. The differential equation for dA/dt (where A is the response) is \frac{d A}{d t}=k_1 \operatorname{S} \cdot(1-A)-k_2 A \cdot B The equation for dB/dt (the inhibitor generated from A) is \frac{d B}{d t}=k_3 \text { S } \frac{1-B}{K_3+1-B}-k_4 B The constants for the graph (right) produced by the Vcell model are: • k1 = 10; k2 = 100 • k3 = 0.1; k4 = 1 • K3 = 0.001 State-dependent Inactivations systems. There are two simple circuits/motifs in this system that were found after the initial analyses that showed all possible interactions in a 3-component system (see Figure 3).  The motif was patterned after the inhibition of proteins in neuron stimulation, specifically in ion channels in neural cell membranes that open up on a change in the transmembrane potential but then close again quickly to avoid constant neuronal stimulation (or inhibition).  In the Naion channel, there are both fast (1-2 ms) and slow (100 ms) inactivation mechanisms.  The fast one allows for repetitive firing, the development of action potentials, and the control of the excitation of neurons, and at the neuromuscular junction.  Neuronal signaling is discussed in Chapter 28.9.  Figure $8$ below shows a simplified model for one type of inactivation of the Naion channel Figure $7$: Simplified state transition model of voltage-gated sodium channels featuring closed, open, and inactivated states.  Zybura, A. et al. Cells 202110, 1595. https://doi.org/10.3390/cells10071595.   Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/). The figure implies that there are at least 3 conformational states of the channel so the inactivation for the channel and the circuit/motif for adaptation we will now discuss are called state-dependent inactivations.  The slow return to the original state is observed in many ion channels as well as in the return of G protein-coupled receptors to the normal state after their desensitization.  Also, some protein kinases (kinases that use ATP to phosphorylate protein substrates) can be inactivated by internalizing the membrane kinase into vesicles where they can be reactivated and returned to the plasma membrane in a slow process. For the construction of a perfect or near-perfect adaption state, we will assume the protein A exists in an off state (Aoff) which binds the stimulus (B or S), an on state (Aon) which is viewed as the response (or A produces the response), and an inactivated state (Ain) which slowly reverts to the Aoff state which can be activated again.  The inactive state can be produced by conformational transitions with the protein itself or another molecule produced downstream of it in a metabolic or signaling pathway. For example, a GPRC could be phosphorylated or bind to another species to produce an inactive state. There are two different circuits/motifs that can produce state-dependent inactivation. We'll refer to these as Type A and Type B Type A Figure $9$ shows the Vcell reaction diagram (top left), a classical reaction diagram (bottom left) and time course graphs for Type A state-dependent inactivation. Figure $9$:  Perfect Adaptation for Type A State-Dependent Inactivation. Adapted from Ferrell (ibid). Aon represents the active state of the protein.  This mechanism applies well to the Na+ channel. The differential equations for dAon/dt and dAoff/dt are shown below. For dAon/dt \frac{d A_{o n}}{d t}=k_1 \operatorname{Input} \cdot\left(1-A_{o n}-A_{i n}\right)-k_2 A_{o n} dAin/dt \frac{d A_{i n}}{d t}=k_2 A_{o n} with constants k1 = k2 = 1. Again, as with the other cases, the stimulus S is pulsed.  The different colors in the bottom left reaction diagram imply an off and inactive red state and a green active state, each of different conformations. The graphs were produced using Vcell.  There is a slight anomaly in the graph of Aon which shows two additional small peaks as the system returns to the basal state.  This contrasts to just 1 peak which returns to the basal state in a simple exponential fashion as described in the Ferrell paper.  We are uncertain as to the source of the discrepancy. Type B In this case, the periodic stimulus, abbreviated as B, is a binding partner for Aoff which produces an active complex B-Aon.   Figure $10$ below shows the Vcell reaction diagram (top left), a classical reaction diagram (bottom left), and time course graphs for Type B state-dependent inactivation Figure $10$ Figure $10$:  Perfect Adaptation for Type B State-Dependent Inactivation. Adapted from Ferrell (ibid) BAon represents the active state of the protein bound to B while BAin represents the inactive complex. The equation of dBAon/dt for the formation of the active state is \frac{d B A_{o n}}{d t}=k_1\left(B_{t o t}-B A_{o n}-B A_{i n}\right) *\left(1-B A_{o n}-B A_{i n}\right)-k_2 B A_{o n} and the equation for dBAin/dt for the formation of the inactive state is \frac{d A_{i n}}{d t}=k_2 A B_{o n} with constants k1 = k2 = 4. The graphs (note the different time concentration scales on the left) show a fairly quick return to the basal state after each pulse of stimuli (B).

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/14%3A_Principles_of_Metabolic_Regulation/14.5%3A_Metabolism_and_Signaling%3A__The_Steady_State_Adaptation_and_Home.txt

• 15.1: Insulin Signaling in the Liver In this section, we will discuss insulin signaling and glycogen synthesis. • 15.2: Glycogenesis The process of forming glycogen is called glycogenesis and it requires the activity of six enzymes. Some of these, we have already discussed including the hexokinase that phosphorylates the 6'-OH of glucose and phosphoglucomutase that converts glucose-6-phosphate to the glucose-1-phosphate isomer. In this section, we will discuss the remaining four enzymes and their role in glycogen biosynthesis. They are Glycogen Synthase, UDP-Glucose Pyrophosphorylase, Glycogenin, & Glycogen Branching Enzyme. • 15.3: Glycogenolyis and its Regulation by Glucagon and Epinephrine Signaling In the previous section, you learned that glucagon signaling down regulates glycogen synthesis. Now lets look at glycogen breakdown, called glycogenolysis and its control by two hormones, glucagon and epinephrine. Only two enzymes are required for the breakdown of glycogen, the glycogen phosphorylase enzyme, and the glycogen debranching enzyme. • 15.4: Regulation of Glycolysis There are three major enzymatic control points within the glycolytic pathway: the hexokinase, the phosphofructokinase, and the pyruvate kinase transitions. Key drivers for regulating the pathway are energy demand within the cell as determined by local indicators such as ATP and AMP, as well as energy demand within the organism as a whole, which can be influenced by hormone signaling pathways. We will also see that the regulation of the pathway can vary depending on cell type and cellular needs. • 15.5: Regulation of Gluconeogenesis There are three major enzymatic control points within the glycolytic pathway: the hexokinase, the phosphofructokinase, and the pyruvate kinase transitions. Key drivers for regulating the pathway are energy demand within the cell as determined by local indicators such as ATP and AMP, as well as energy demand within the organism as a whole, which can be influenced by hormone signaling pathways. We will also see that the regulation of the pathway can vary depending on cell type and cellular needs. 15: Glucose Glycogen and Their Metabolic Regulation Search Fundamentals of Biochemistry Introduction In this section, we will discuss insulin signaling and glycogen synthesis. Insulin is released in the fed state, and leads to glucose uptake where it can be stored, if not needed, for glycogen synthesis. Recall that glycogen is a large polymer of glucose residues connected in the main chain by alpha 1 → 4 linkages and with branching side chains about every 12 – 15 residues at the alpha 1 → 6 positions. The reducing ends of the carbohydrate (two for each polymer) are connected to the glycogenin dimeric protein at the center of the macromolecule (Figure \(1\)). With the branching nature of the polymer, many non-reducing ends of the molecule are present, allowing easy access and fast release of glucose for energy utilization. Most of the body’s pools of glycogen are stored in the liver, with 10% of the liver biomass in glycogen granules, and in the skeletal muscle, with glycogen comprising 2% of the biomass of the muscle. Each glycogen polymer may have upwards of 30,000 glucose residues, making the glycogen polymer visible using standard microscopic techniques. Storage of glycogen within muscle tissue is used by the muscle cells as a source of energy to fuel muscle contraction. In the liver, the purpose of glycogen storage is different. Glycogen stored at this location is used to maintain the homeostatic balance of blood glucose levels. The liver is the primary organ that can actively transport glucose into the bloodstream. Our only other major source of glucose within the blood is our diet. Blood glucose homeostasis is critical for brain function (Figure \(2\)). The brain has a huge energy demand, but nearly zero storage of key energy molecules required for ATP production. Furthermore, glucose and ketone bodies are the only energy sources that can pass the blood-brain barrier and be utilized by the brain for ATP production. Note that ketone bodies are only produced during starvation, or disease states such as diabetes, and are not a regular source of energy for the brain. Thus, glucose is critical for brain function. Nearly 10% of the whole body’s energy is used for nerve impulse transmission by the brain. If blood flow to the brain carrying critical oxygen and glucose is impeded, people will lose consciousness within approximately 20 seconds! And brain death/permanent damage occurs within 4 minutes of blood flow cessation. This exemplifies the importance of the liver in maintaining blood glucose levels, as well as the importance of oxygen maintenance. Glycogen in the liver or muscle can be broken down into glucose 1-phosphate (Figure \(3\)). This can be interconverted to glucose 6-phosphate which is then readily used in many cellular processes. The process of glycolysis (or the breakdown of glucose into pyruvate) occurs in all cells and produces a small amount of ATP in the process. Further processing of pyruvate can occur anaerobically (or in the absence of oxygen) to produce lactate, or the process can continue to occur in the aerobic pathway to complete oxidation to carbon dioxide and water in the Kreb cycle. Note that oxygen from breathing is used to create the water within this pathway. This fuels the process of oxidative phosphorylation within the mitochondria and produces large quantities of ATP (from 30-36 molecules/glucose). Within the liver, glucose can be freed from glycogen and released back into the bloodstream to maintain homeostatic levels. Glucose 6-phosphate can also be utilized as a precursor for other major macromolecules such as ribose and deoxyribose, as well as the hexosamine compounds commonly found cushioning joints or attached to proteins of the plasma membrane. In healthy individuals, hormone signaling is critical to maintaining blood glucose homeostasis. Within this system, the hormones glucagon and insulin work together to maintain normal plasma glucose levels ( Figure \(4\)). During hyperglycemia, pancreatic beta (β) cells release insulin, which stimulates glucose uptake by energy-consuming cells and the formation of glycogen in the liver. During hypoglycemia, pancreatic alpha (α) cells release glucagon, which stimulates gluconeogenesis and glycogenolysis in the liver and the release of glucose to the plasma. The first area we will focus our attention on will be the mechanism utilized by insulin to reduce blood glucose levels. Figure \(5\) shows the structure of the pancreas and its anatomical relationship with the liver and the stomach. The pancreas is the sensor organ that detects blood glucose levels. It is responsible for signaling to the liver to either remove or release glucose in response to changing levels. Notably, the pancreas also produces most of the digestive enzymes utilized by the body, including proteases, amylases, and lipases. Figure \(6\): shows a light microscope image of the pancreatic islet cells. They are responsible for the production of glucagon and insulin. The islets are distinguished from the surrounding tissue by a continuous connective tissue capsule and extensive vascularity. Insulin is a peptide hormone composed of 51 amino acids as shown in Figure \(7\). It is initially synthesized as preproinsulin, which is converted to proinsulin after the signal peptide is removed.  Two disulfides are made in the ER (catalyzed by protein disulfide isomerase) along with selective proteolytic cleavage to form insulin.  Mature insulin consists of A and B chains that were connected in proinsulin by the C-peptide. Figure \(7\): Conversion of preproinsulin to mature insulin.  Vasiljević, J. et al. Diabetologia 63, 1981–1989 (2020). https://doi.org/10.1007/s00125-020-05192-7.  http://creativecommons.org/licenses/by/4.0/. Figure \(8\) shows  interactive iCn3D models of human insulin (3I40) and human proinsulin (2KQP).  (Copyright; author via source). Click the image for a popup or use the external link provided: Human insulin (3I40) Human Proinsulin (2KQP) Colored code to show secondary structure External link:  https://structure.ncbi.nlm.nih.gov/i...A3QZdZpfEMxR17 The "future" A chain in mature insulin is shown in magenta, the C-peptide connecting the A and B chains is shown in yellow, and the future B chain is in cyan. External link:  https://structure.ncbi.nlm.nih.gov/i...nQJXhrEfHD1GH9 The maturation of preproinsulin to insulin is shown in more detail in  Figure \(9\).  The peptide is first translated on ribosomes linked to the rough endoplasmic reticulum (ER), where a signal peptide docks the peptide to the ER membrane. The proinsulin is folded and the signal peptide is cleaved. It is transported to the Golgi where it is further packaged into secretory vesicles. Within the secretory vesicles, the proinsulin is cleaved to release the C-peptide. The A and B peptides are held together by disulfide bridges and form the active insulin component. The C-peptide is a bioactive peptide secreted at the same time and in equimolar amounts to the insulin hormone. It also has a longer half-life than insulin and is excreted by the kidneys into the urine, making detection easy. Furthermore, it allows for the detection of patient-produced insulin, even if they are receiving insulin injections. Thus, C-peptide detection is often utilized to help distinguish between patients with type 1 diabetes from patients with type 2 diabetes (or Maturity onset diabetes). Details about the different forms of diabetes will be discussed in greater detail later. Once insulin is released from the pancreas, it travels throughout the body and binds with cellular targets that contain the insulin receptor. The Insulin Receptor is a tyrosine kinase receptor that dimerizes upon insulin binding, as shown in Figure \(10\). Insulin receptors are located on most cell types throughout the body causing pleiotropic effects during insulin response. Primary targets of insulin action are the liver, where it promotes the uptake of glucose and the production of the glycogen storage molecule, as well as skeletal muscle and fat. The tyrosine kinase portion of the receptor located on the internal side of the plasma membrane is quite flexible. The lefthand diagram shows a space-filling model of the activated insulin receptor dimer embedded into the plasma membrane (shown as the gray bar). The tyrosine kinase portion of the receptor is shown on the inside of the cell whereas the insulin binding domain is present on the external side of the plasma membrane. The middle and right-hand diagrams show the inactive (middle) and active forms (far right) of the tyrosine kinase domain of an insulin receptor monomer. When activated the tyrosine kinase domain binds to ATP (hot pink) and phosphorylates downstream targets, including several of its tyrosine residues (green). In the inactive state (middle), a mobile loop (turquoise) binds in the ATP binding site and prevents ATP association. When the insulin receptor is activated, the mobile loop opens, allowing for the binding of ATP and self-phosphorylation of tyrosine residues, as well as other signaling proteins (a small peptide from one is shown in light pink). Figure \(11\) below which shows an interactive iCn3D model of the Full-length mouse insulin receptor bound to four insulins (7SL7). Figure \(11\): Full-length mouse insulin receptor bound to four insulins (7SL7). (Copyright; author via source). Click the image for a popup or use this external link:  https://structure.ncbi.nlm.nih.gov/i...gkTrtvaqQF6o2A The receptor is a dimer (one monomer gray and the other light brown).  Four insulins are bound in maximally activated insulin receptors.  Two insulins (magenta) are bound at the same respective place in each monomer (site-1, the primary site) and the two others are bound at a second parallel site (site-2).  The full active state is a symmetric T-shape.  Less active receptors have fewer bound insulins with receptor geometry more asymmetric (one insulin bound at site 1 gives a Γ-shaped conformation, while two produce a Ƭ-shaped conformation as the second insulin binds). When 4 insulins are bound at both sites, the asymmetric conformation can't be formed.  Although the structure is described as full-length, both monomers end at amino acid 910.  The single membrane-spanning alpha-helix membrane occurs at amino acids 947-967 and is NOT shown in the model. Activation of the insulin receptor in the liver when insulin is present initiates a phosphorylation signaling cascade, as shown in Figure \(12\).  One function of the signaling cascade results in the activation of the Rab10 protein. Rab 10 promotes the fusion of GLUT4-containing secretory vesicles (GSVs) with the plasma membrane allowing for increased surface expression of GLUT4. GLUT4 is a glucose transporter protein. Thus, an increased concentration of the protein in the plasma membrane results in the upregulation of glucose import into the cell. Having GLUT4 proteins stored within secretory vesicles makes it available more readily than having to activate gene transcription pathways and production of the protein de novo. This allows a faster response to help lower blood glucose levels. The result is increased glucose uptake from the bloodstream into liver cells and other cellular targets, reducing blood glucose levels. The insulin receptor is a receptor tyrosine kinase, which undergoes dimerization and autophosphorylation of Tyr residues upon insulin binding. The phosphorylated receptor also recruits and phosphorylates the insulin receptor substrate 1 (IRS-1) on tyrosine residues, which then recruits dimeric Phosphoinositol (PI)3-kinase (p85/p110 in the diagram above) and phosphorylates the p85 regulatory subunit. The PI3 kinase catalyzes the phosphorylation of phosphatidylinositol bisphosphate (PIP2) within the plasma membrane to form phosphoinositol, 3,4,5-triphosphate (PIP3). PIP3 then recruits PIP3-dependent kinase (PDK) which phosphorylates and activates Akt. Once activated, Akt dissociates from the membrane into the cytosol where one of its downstream targets is AS160. AS160 is a GTPase that normally binds with Rab10 (a G-protein) causing the cleavage of GTP to GDP. Thus, AS160 downregulates the activity of Rab10. In the phosphorylated state, AS160 cannot bind or inhibit Rab10, enabling Rab10 to release GDP and bind with a molecule of GTP. In the activated state, Rab10 helps promote the fusion of GLUT4-containing secretory vesicles (GSVs) secretory vesicles with the plasma membrane. Figure \(13\) provides a deeper look at some of the initial activation steps in the insulin signaling pathway. This step shows the phosphorylation of Phosphoinositol 4,5-bisphosphate (PIP2) to Phosphatidylinositol 3,4,5-triphosphate (PIP3). PIP2 is a common phospholipid within the lipid bilayer structure. In future lectures, we will see the utilization of this phospholipid in other signaling pathways as well. Once glucose enters a cell, it is rapidly converted to glucose 6-phosphate via the enzyme hexokinase, as shown in Figure \(14\) below. This enzyme is covered in more detail in our section on glycolysis. Importantly, phosphorylation traps the glucose inside the cell and does not allow it to be redistributed back into the bloodstream. This helps to maintain the homeostasis of glucose within the bloodstream. In addition, glucose 6-phosphate is the first step in many pathways utilizing glucose, including energy utilization and the formation of building blocks such as ribose and deoxyribose used in RNA and DNA synthesis. Insulin signaling increases the number of GLUT4 transporters in the plasma membrane causing an increased uptake of glucose into the cell. Within liver and muscle tissue, if glucose is not required for energy or other metabolic intermediates, it is then converted to glycogen for storage. The major enzyme required for glycogen synthesis is also activated via insulin signaling, as shown in Figure \(15\).  In addition to phosphorylating the AS160 protein, Activated Akt also phosphorylates the Glycogen Synthase Kinase enzyme (GSK-3) which inactivates this protein. This allows protein phosphorylase 1 (PP1) to dephosphorylate the Glycogen Synthase enzyme shifting it into a more active state, and causing glycogen synthesis to commence. In this section, we have covered two pathways activated during cellular response to insulin signaling, and I am sure that you are feeling a bit overwhelmed with the complexity. However, biological processes are incredibly complex and signaling pathways have multiple pleiotropic downstream effects. For insulin signaling, we have only touched the tip of the iceberg, as evidenced in Figure \(16\). This figure gives a more complete representation of the chemical changes induced within a liver cell in response to insulin signaling. For our purposes, we will restrict coverage to the two downstream effects: an increase in GLUT4 transporters in the plasma membrane and increased activity of glycogen synthase.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/15%3A_Glucose_Glycogen_and_Their_Metabolic_Regulation/15.01%3A_Insulin_Signaling_in_the_Liver.txt

Search Fundamentals of Biochemistry Introduction The process of forming glycogen is called glycogenesis and it requires the activity of six enzymes as illustrated in Figure $1$.  We have already discussed several including hexokinase which phosphorylates the 6'-OH of glucose and phosphoglucomutase which converts glucose-6-phosphate to the glucose-1-phosphate isomer. In this section, we will discuss the remaining four enzymes and their role in glycogen biosynthesis. They are Glycogen Synthase, UDP-Glucose Pyrophosphorylase (preferred name UTP-glucose-1-phosphate uridylyltransferase), Glycogenin, and Glycogen Branching Enzyme. Given the importance of these enzymes in the synthesis of the second main energy storage molecule, we must probe the enzymes in detail. Glycogen synthase (GS) is a key enzyme and its activity is highly regulated.  In Chapter 15.1, we have already explored how insulin signaling upregulates the activity of this enzyme by inhibiting phosphorylation by GSK-3. Other effectors include the allosteric binding of glucose-6-phosphate, which also increases the activity of the GS. In a later section, we will also see that the hormone glucagon can also regulate the activity of the GS through protein kinase A (PKA) in a fashion that decreases glycogen synthesis and increases glycogen breakdown. In the glycogenesis pathway, GS is responsible for building the majority of the main alpha 1 → 4 chain glucose acetal linkages. The GS does require a primer of 4 -6 glucose residues linked together by alpha 1 → 4 bonds to begin synthesis. Since GS can only form alpha 1 → 4 linkages in the main chain, it CANNOT create the alpha 1 --> 6 branches inherent to the core structure of glycogen. To build the glycogen main chain, GS uses the glycogen primer and glucose that has been activated through covalent attachment to uridine diphosphate (UDP) at the 1-position. Upon completion of one round of synthesis, the 1 position of the incoming UDP-glucose is covalently attached to the 4-position of the nascent glycogen molecule, releasing the UDP as a leaving group. $\text { Glycogen }_{(n)}+\text { UDP-glucose } \rightarrow \text { Glycogen }_{(n+1)}+\text { UDP } \nonumber$ UTP--glucose-1-phosphate uridylyltransferase (or UDP-Glucose Pyrophosphorylase) The formation of the UDP-glucose required for the synthesis of the main chain of glycogen is mediated by UTP-glucose-1-phosphate uridylyltransferase (preferred name), which is also called UDP-glucose pyrophosphorylase (GalU or UGPase; EC 2.7.7.9). UGPase catalyzes the reversible reaction of glucose 1-phosphate and UTP into UDP-glucose and inorganic pyrophosphate (PPi) (Figure $2$). Enzymes of the UGPase family are ubiquitous and can be found in the tree of life. UDP-glucose is an activated form of glucose used in the synthesis of other glycans including sucrose, cellulose, start, and glycogen, and the glycan parts of glycoproteins, glycolipids, and proteoglycans.  Hence it is a key metabolite, which gives more importance to understanding UGPase. Like many other nucleotidyl transferases, UGPase requires divalent cations to promote the reaction (Figure $3$). In most cases, magnesium ions are employed. The reaction mechanism follows a sequential bi-bi-mechanism starting with the binding of UTP to the active site, in presence of a magnesium ion, followed by the binding of glucose 1-phosphate. The octahedral coordination sphere of the magnesium positions the substrates in the right way and enables the nucleophilic attack of glucose 1-phosphate on UTP. A lysine, an aspartate, and several water molecules within the active site help to stabilize the position of the substrates and cofactor for the proper nucleophilic attack of the phosphoryl oxygen of glucose 1-phosphate towards the α-phosphorous atom of UTP. Finally, PPi is released from the UGPase/Mg2+/UDP-glucose complex. UDP-Glucose then dissociates from the complex restoring the active site of the enzyme for another round of synthesis. Figure $4$ shows an interactive iCn3D model of the human UDP-glucose pyrophosphorylase tetramer (3R2W). Figure $4$:  Human UDP-glucose pyrophosphorylase tetramer (3R2W). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?9yq9STPGvq87Wnbt8 Substrate binding and active site residues are shown as CPK-colored sticks and labeled in one of the subunits.  Three key loops whose correct positioning is required for catalysis are colored as follows: • Latch Loop:  406-416, yellow, which contains Glu 412 (shown in spacefill) • SB Loop:  275-282, red • 309 Loop: 309-311, cyan, which contains Ser 309 (shown in spacefill). Only four of the eight subunits are shown for clarity.  Note the proximity of the interacting loops positioned between the brown and magenta subunits.  The SB and 309 loops at that location are part of the brown subunit, while the yellow latch loop is part of the magenta subunit.  The latch loop is between the SB and 309 loops.   The cyan 309 loop hence is prevented from interacting with the substrate and with the movements of the SB and 309 loops needed for catalysis.  In other species, it has been observed the SB loop moves down UDP-glucose on binding to the active site.  The structure above is for the apo-enzyme without bound substrate, and hence the inactive or closed form of the protein. Mutations in the 309 loop (S309N/S311R) still had 84% of normal activity.  Glu412 in the latch loop is highly conserved in vertebrates (but not present in yeast).  Mutations in Glu 412 didn't affect the formation of oligomers of the enzyme but did affect activity. • replacing E412 with a short aspartate (E412D) significantly increase activity  (176%) • E412Q, which eliminates the charge while retaining the approximate size of the side chain showed just a marginal increase in activity (19%) • E412K, which flips the charge and increases the length of the side chain decreased activity to 22%. These mutations generally suggest that steric effects in the region of subunit interaction are most important in activity. Glycogen Synthase UDP-glucose is then utilized by glycogen synthase (GS) to extend the main chain of glycogen by one glucose residue. In this reaction, the 4’-OH group of the glycogen main chain attacks the anomeric carbon of UDP-glucose (Figure $5$). The UDP functional group serves as a good leaving group allowing for the formation of the alpha 1 --> 4 bond. Glycogen synthase has two isoforms, GS1 expressed in tissue where glycogen is present (for example muscle) and GS2 expressed only in the liver.  We will explore its mechanism more fully below. Glycogenin Previously, we mentioned that GS requires a glycogen primer of 4 – 6 glucose residues to begin adding new residues to the main chain. This primer is provided by the small docking protein, Glycogenin (GN or GYG). This protein is a homodimer that self-catalyzes its own glycosylation at amino acid Tyr-194. In this reaction, UDP-glucose is coordinated by a Mn2+ metal cofactor and critical aspartate residues (Figure $6$). The –OH group of Tyr-194 then mediates nucleophilic attack on the anomeric carbon of UDP-glucose. Thus, glycogenin is tethered to the reducing end of the glycogen molecule. As with glycogen synthase, glycogenin (GN or GYG) has two isoforms, with GN2 (or GYG2) expressed mostly in the liver, pancreas, and heart. Mechanisms for glycogen synthesis by glycogenin and glycogen synthase How do glycogenin and glycogen synthase cooperate in the synthesis of glycogen?  Structures of the complex of glycogenin-1 (GYG1), which seeds the molecule by starting glycogen synthesis by autoglucosylation, and glycogen synthase-1 (GS1 or GYS1), which extends the molecule, show allosteric transitions between three main states, the closed/inactive, partially open, and open active complex, as shown in Figure $7$: Figure $7$: GYS1 chains A, B, C, and D are colored orange, turquoise, purple, and navy respectively. GYG1 globular domains and the GYG1-tail fragment are colored in gray shades for all chains.  The left bottom structure shows the apo GYS1:GYG1 mobile complex.  The bottom middle structure is the apo GYS1:GYG1 ordered complex.  The bottom right structure is the +G6P GYG1:GYS1 complex.  Fastman et al., 2022, Cell Reports 40, 111041 July 5, 2022, 2022. https://doi.org/10.1016/j.celrep.2022.111041.  Creative Commons Attribution (CC BY 4.0) It makes sense that both enzymes bind to each other and cooperate in the synthesis of glycogen. In the closed state, GS (GYS) is a tetramer, which like hemoglobin can be described as a T or closed/inactive state.  In a slight difference, one of the GS (GYS) subunits appears to display an asymmetric conformation which leads to close interactions with GN (GYG) allowing the glycogen seed polymer on GN (GYG) to move to GS (GYS) for elongation (a partially open state).  Multiple conformations of the complex have been resolved.  Further conformational changes lead to the open/active state and a more open binding groove for GN (GYG). Figure $8$ shows an interactive iCn3D model of the  Human glycogenin-1 and glycogen synthase-1 complex in the presence of glucose-6-phosphate (8CVX).  Glucose-6-phosphate is an allosteric activator of glycogen synthase. Figure $8$:  Human glycogenin-1 and glycogen synthase-1 complex in the presence of glucose-6-phosphate (8CVX). (Copyright; author via source). Click the image for a popup or use this external link:  https://structure.ncbi.nlm.nih.gov/i...ivh4gZCupGcXZ7 The larger glycogen synthase subunits in the tetramer are shown in various colors while the glycogenin fragments bound to each monomer of GS (GYS) are shown in gray. Figure $9$ illustrate in greater detail the conformational changes induced on the binding glucose-6-phosphate to glycogen synthase. Figure $9$: G6P binding induces a conformational change across the GYS1 tetramer leading to an open conformation for all four active sites.  Fastman et al, ibid. Panel (A) shows a cartoon representation of the +G6P GYS1:GYG1 complex (left) with GYS1 and GYG1 chains colored as in the previous figure. G6P is shown as spheres colored by heteroatom. Blue and yellow lines and boxes (right) indicate relative positions for perpendicular views. A black dotted circle indicates the oligomeric interface between the CTD-loop region and tetramerization core domains. The CTD-loop, residues 484–488, forms cross-protomer interactions on the other side of the active site.  Green dashed semicircles indicate open active sites. Panel (B) shows G6P-binding site interactions. G6P (white) and proximal interacting residues are shown as sticks and colored by heteroatom. Polar interactions are indicated by yellow dotted lines. Panel (C) shows changes at the G6P-binding site across conformations. A G6P-bound protomer is highlighted in orange with G6P shown as spheres. The adjacent active state protomer (chain C) is shown as a green cartoon. A basal state protomer (chain C) is modeled relative to the orange G6P-bound protomer and shown in yellow. Key changes to the G6P-sensing loop and a regulatory helix are highlighted by outline and non-transparent representation with arrows indicating the relevant motions. The rest of each chain is shown in a transparent cartoon representation. We will return to the mechanism after exploring the last enzyme in the pathway. Glycogen Branching Enzyme The final enzyme, the glycogen branching enzyme (GBE), catalyzes the hydrolytic cleavage of an α(1→4) glycosidic linkage and subsequent inter- or intra-chain transfer of the non-reducing terminal fragment to the C6 hydroxyl position of an α-glucan (Figure $10$). In this example, an inter-chain transfer is occurring. At the top of the scheme, above the arrow, you can see that the GBE enzyme transiently removes several glucose residues (usually around 7) from one linear glycogen chain and then attaches it as an alpha 1→ 6 branch to the other chain. In this process, an additional non-reducing end is created which can act as a primer site for Glycogen Phosphorylase (the main enzyme that breaks down glycogen). Thus, glucose residues can be released very quickly when needed. Details of the structure and domain organization of the human glycogen branching enzyme are shown in Figure $11$. The four domains include N1, CBM48, the catalytic domain, and the C-terminal domain. Figure $11$:  Crystal structure of hGBE1.   Froese et al. Human Molecular Genetics, Volume 24, Issue 20, 15 October 2015, Pages 5667–5676, https://doi.org/10.1093/hmg/ddv280.  Creative Commons Attribution License (http://creativecommons.org/licenses/by/4.0/) Panels (A and B) show orthogonal (perpendicular) views of hGBE1 showing the N-terminal helical segment (orange), CBM48 (pink), central catalytic domain (green), and C-terminal domain (blue). The catalytic triad Asp357-Glu412-Asp481 is shown as red sticks. Numbers refer to domain boundaries. N- and C-termini are labeled as grey spheres. Panel (C) shows the superposition of branching enzyme structures from human (hGBE1, this study), O. sativa SBE1, and M. tuberculosis GBE, highlighting the conserved domain architecture and three regions of structural variation. Panel (D) shows the domain organization of hGBE1, O. sativa SBE1, and M. tuberculosis GBE revealing differences in the N-terminus between prokaryotic and eukaryotic polypeptides. Prokaryotic GBEs contain two N-terminal carbohydrate-binding domains (N1, N2) whereas eukaryotes contain only one (CBM48) and replace the prokaryotic N1 domain with a helical extension. Figure $12$ shows an interactive iCn3D model of the human glycogen branching enzyme (GBE1) Figure $12$: human glycogen branching enzyme (GBE1). (Copyright; author via source). Click the image for a popup or use this external link:  https://structure.ncbi.nlm.nih.gov/i...47aqw5qto8hkU7 The domains in the iCn3D module are colored similarly to that of the previous figure. The enzyme core is similar to amylase with the conserved active site.  Diseases of glycogen storage often result from mutations in the amylase core domain. For example, late-onset adult polyglucosan body disease (APBD) arises from a common mutation, Y329S.  The effect of this mutation may arise from misfolding.  A tetrapeptide, Leu-Thr-Lys-Glu, given to patients, increased activity twofold, probably by acting as a chaperone to facilitate proper folding. Putting it all together: Glycogen Synthesis and Its Regulation Figure $13$ shows the steps involved in the addition of glucose from the donor UDP-glucose to glycogenin and through glycogen synthase to the growing glycogen polymer which becomes branched through the activity of the glycogen branching enzyme. Figure $13$:  Summary of glycogen synthesis. Marr, L., Biswas, D., Daly, L.A. et al. Nat Commun 13, 3372 (2022). https://doi.org/10.1038/s41467-022-31109-6.  Creative Commons Attribution 4.0 International License.  http://creativecommons.org/licenses/by/4.0/ The top reaction shows the step catalyzed by glycogenin (GN), while the bottom reactions are those catalyzed by glycogen synthase and the glycogen branching enzyme. Regulation of glycogen synthesis occurs in part through the phosphorylation of key residues in both GS (GYS) and GN (GYG).  Those sites are shown in Figure $14$. Figure $14$: Domain and phosphorylation sites for glycogen synthase 1 (GS1) and glycogenin 1 (GN1).  Marr et al, ibid Panel c shows the domain architecture of human GS (top) and GN (bottom). Known in vivo phosphorylation sites of GS are shown in red and are labeled with residue number and classical nomenclature (in bold). GN tyrosine 195 that becomes auto-glucosylated and was mutated to phenylalanine (Y195F) in this study is indicated. Not to scale. Panel e shows the cartoon representation of GN WT and Y195F. As we showed above, glycogen synthase is allosterically activated on the binding of glucose-6-phosphate, which can be thought to activate the enzyme by a T to R state change.  The enzyme is inactivated by phosphorylation at multiple sites as shown above. Activation hence can also occur through dephosphorylation.  Phosphorylation of a site can provide a binding site that leads to more phosphorylations.  This can lead to a flexible  "spike" of hyperphosphorylated residues forming from two of the monomers. Particularly important is pSer641 (site 3A) which interacts with a series of arginine residues in a regulatory helix of glycogen synthase.  This arginine cluster has been called the arginine cradle.  The interaction sites are illustrated in Figure $15$. Figure $15$: The phosphoregulatory region of human GS. Marr et al, ibid Panel a shows the human (Hs)GS-GN34 structure in ribbons (top left). The N- and C- terminal tails of one GS protomer (chain A) lie next to one another and move towards the adjacent protomer, meeting the N- and C-terminal tails from chain B. Arrows indicate a continuation of cryo-EM density (top right). Electron density (C1 symmetry) for phosphorylated S641 (pS641) interacting with R588 and R591 on the regulatory helices α22 (bottom left). Residues that are interacting with the N- and C-terminal tails that are mutated in this study are shown (bottom right). Panel b shows a comparison of distances between regulatory helices of adjacent monomers of HsGS (reported here), low activity inhibited mimic (PDB ID 5SUL), basal state (PDB ID 3NAZ), and G6P activated (PDB ID 5SUK) yeast GS (yGS) crystal structures. Quoted distances were measured from Cα of Arg591 (chain A) and -Cα of Arg580 (chain B) of HsGS and corresponding yeast residues. The strong electrostatic arginine-pSer interactions lock the tetramer into the inactive T state. A cartoon model illustrating the regulation of glycogen synthase by phosphorylation/dephosphorylation and interconversion between T and R state is shown in Figure $16$. Figure $16$: GS and GN cooperate to synthesize glycogen. Marr et al, ibid The inhibition by phosphorylation can be relieved by binding the allosteric effector glucose-6-phosphate and does not require phosphatases,

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/15%3A_Glucose_Glycogen_and_Their_Metabolic_Regulation/15.02%3A_Glycogenesis.txt

Search Fundamentals of Biochemistry In the previous section, you learned that glucagon signaling down-regulates glycogen synthesis. Now let's look at glycogen breakdown, called glycogenolysis, and its control by two hormones, glucagon, and epinephrine. Only two enzymes are required for the breakdown of glycogen, the glycogen phosphorylase enzyme, and the glycogen debranching enzyme. Glycogenolysis: An Overview Two key enzymes are required for the stepwise catabolism of glycogen, glycogen phosphorylase, and glycogen debranching enzyme. In the liver, the ultimate end product is glucose-1-phosphate, which is dephosphorylated in the liver to enable the export of free glucose into the circulation. In contrast, in the respiring skeletal muscle, it is converted to glucose-6-phosphate for use in glycolysis. Glycogenolysis is also activated by the hormones glucagon and epinephrine. Glycogen Phosphorylase Glycogen phosphorylase (GP) catalyzes the release of glucose 1-phosphate from the alpha 1→ 4 non-reducing ends of glycogen. An overview of this reaction is shown in Figure \(1\). Glycogen phosphorylase is a homodimer with two active sites. It also requires a cofactor, pyridoxal phosphate (PLP) to be functional (Figure \(2\)). The PLP is derived from Vitamin B6. You may have heard previously that low levels of  B vitamins are associated with lethargy or a lack of energy. We will continue to see that the B vitamins provide essential cofactors for enzymes involved in the production of ATP. Thus, if you lack B vitamins, you are not efficiently producing ATP. The PLP cofactor of GP is attached covalently to the enzyme through a Schiff-base linkage with a Lysine (K) residue. The reaction mechanism of glycogen phosphorylase is detailed in Figure \(3\). When glycogen phosphorylase binds with glycogen a free inorganic phosphate anion is positioned by the PLP and the enzyme active site in proximity with the anomeric carbon position of the non-reducing end residue of the glycogen molecule. The oxygen involved in the glycosidic bond attacks the partially charged hydrogen associated with the phosphate ion, leading to the cleavage of the glycosidic bond. The cleaved glycogen chain leaves the active site and one of the phosphate oxygens attacks the carbocation intermediate created during the cleavage. This results in the release of the terminal glucose residue as glucose 1-phosphate Glycogen Debranching Enzyme Glycogen phosphorylase cannot cleave the alpha 1 → 6 linkages, and it also cannot cleave alpha 1 --> 4 linkages that are within 4 residues of an alpha 1→ 6 linkage (the glycogen chain will no longer fit into the active site of the enzyme) The Glycogen Debranching Enzyme (GDE) has two catalytic activities that enable it to deal with this problem. The first catalytic activity is a Glycosyl Transferase (GT) activity. In this process, the three remaining alpha 1 → 4 extended units on the branch site (colored in green) are clipped off of the branch site and attached to a straight chain of alpha 1 → 4 extended glucose residues. The second part of the reaction requires the Glucosidase (GC) activity that mediates the hydrolysis of the alpha 1 → 6 glycosidic bond and the release of free glucose in the process. Glycogen Phosphorylase can then resume the breakdown of the remaining alpha 1 → 4 chain. An overview of glycogen breakdown is shown in Figure \(4\). Dephosphorylation of Glucose 1-Phosphate Following the activation of glycogenolysis, the liver cell has now released large quantities of glucose 1-phosphate from glycogen, as well as a smaller amount of free glucose from the clipped branch residues. The free glucose can be transported to the bloodstream straight away, but the glucose 1-phosphate must be dephosphorylated prior to release (Figure \(5\)). The dephosphorylation of glucose only occurs in liver cells, as this is the primary location for the regulation of blood glucose levels. Free glucose can exit the cell while phosphorylated forms are trapped inside the cell. Figure \(6\) outlines the process of glucose dephosphorylation in the liver. To mediate the dephosphorylation of glucose, glucose 6-phosphate is transported from the cytoplasm into the lumen of the endoplasmic reticulum (ER) through transporter 1 (T1). The glucose 6-phosphatase (G-6-Pase) then cleaves the phosphate from the substrate, releasing inorganic phosphate (P) and glucose (red molecule). The inorganic phosphate is then transported back into the cytoplasm through transporter 2 (T2) and glucose is transported through Transporter 3 (T3). Free glucose is then transported back into the bloodstream through a glucose (GLUT) transporter located in the plasma membrane. Hormonal Control of Glycogen breakdown In the previous sections, we’ve discussed insulin signaling and the process of building glycogen (glycogenesis) in detail. Now let’s take a look at the other side of the homeostatic balance which begins with glucagon signaling. During hypoglycemia (or low blood glucose levels), pancreatic alpha (α) cells release the hormone peptide, glucagon, which stimulates gluconeogenesis (the formation of glucose) and glycogenolysis (the breakdown of glycogen) in the liver, resulting in the release of glucose to the plasma, and the raising of blood glucose levels, as shown in Figure \(7\). Let’s review a few terms before we begin. In the last section, we were introduced to glycogenesis, the synthesis of glycogen. We saw that this pathway was activated during insulin signaling. In glucagon signaling, this pathway is inhibited and the opposite pathway, glycogenolysis (glycogen breakdown) is activated. Glucagon signaling in the liver also down-regulates glycolysis (the utilization of glucose for energy production), as the liver is trying to use glucose to maintain blood glucose levels. It doesn’t utilize it for its own energy needs during this time. Instead, lipids can be used by liver cells to generate ATP, and in fact, glucagon signaling increases lipolysis or the breakdown of lipids. Finally, glucagon also up-regulates the process of gluconeogenesis or the generation of glucose from non-sugar metabolites. We will address the mechanisms of glycolysis and gluconeogenesis regulation in a later section. Here we will only take a cursory look at these pathways and will focus more on the process of glycogenolysis. Overview of Glucagon Signaling Glucagon signaling begins when the hormone binds with its receptor on liver cells as shown in Figure \(8\). Glucagon receptors are not widespread within the body as insulin receptors. Since the purpose of this hormone is to cause the release of glucose back into the bloodstream, this process is highly controlled and only the liver can deliver glucose back into the bloodstream to maintain homeostasis. Thus, other target tissues such as skeletal muscle do not need to have these receptors expressed and are not sensitive to glucagon signaling. The glucagon receptor is a G-protein-coupled receptor and is also referred to as a 7TM receptor (as it contains 7 transmembrane domains that span the plasma membrane). This family of receptors is widespread throughout the body and responsible for many of the pharmaceutical mechanisms of action seen in our treatment of different disease conditions. With regards to this pathway, once glucagon binds to the receptor, the receptor moves laterally in the plasma membrane and binds with a G-protein that is stationed as a peripheral protein to the plasma membrane. The G-protein contains three major domains, the alpha, the beta, and the gamma domain. The alpha domain is capable of binding to the GDP/GTP cofactor. When the G-protein is inactive, all three subunits stay together and the alpha subunit remains inactive and bound to GDP. When the G-protein associates with an activated receptor, the alpha subunit exchanges GTP for the bound GDP cofactor, and the gamma and beta subunits dissociate into the cytoplasm. The activated alpha subunit moves laterally on the periphery of the plasma membrane until it contacts the adenylyl cyclase enzyme (also called adenylate cyclase). This activates the adenylyl cyclase that converts ATP into cyclic AMP (cAMP). cAMP production is an amplification step within this pathway. That means that more cAMP is produced than G-proteins are activated. After some time a G-protein hydrolase will cause the hydrolysis of the GTP to GDP and inactivate the G-protein. At this point, the G-protein will associate with the gamma and beta subunits reforming its inactive state. Another glucagon signaling event will be required to reactivate the process. The cyclic AMP produced in the process serves as a second messenger in the process and activates a myriad of downstream targets. We will focus on two of the major targets. The first is Protein Kinase A, which becomes activated upon binding with cAMP. The second target is a cAMP Response Element-Binding Protein (CREB). The CREB protein is also activated when bound to the cAMP molecule. This causes the CREB protein to translocate from the cytosol into the mitochondria and the nucleus. In both of these locations, the activated CREB binds to specific response element sequences in the DNA and activates the transcription of genes that are involved in gluconeogenesis. These genes and their encoded proteins have been discussed in more detail in chapter 14. What is important to note now is that glucagon signaling in the liver results in the upregulation of glucose production de novo from non-carbohydrate precursors. This is NOT a favored pathway in the body. It is expensive energetically for the liver to manufacture glucose. In fact, more expensive in the cost of ATP than can be produced from the newly formed molecule. However, organs like the brain can only utilize free glucose as an energy resource. Thus, the liver will engage in this energy deficit to build glucose for use by the brain and other cellular targets. Glucagon signaling also leads to the downregulation of glycolysis, which we will cover in more depth in section 15.4 and glycogenesis. It also leads to an increase in glycogenolysis or the breakdown of glycogen. Let's take a further look at the regulation of both of these processes. Regulation of Glycogenesis - Since glycogen synthase (GS) is the primary enzyme required for glycogenesis, it is also the primary target for the regulation of this pathway. Recall that GS is active in the dephosphorylated state. Thus, PKA down-regulates the activity of this enzyme through the phosphorylation of GS, as shown in Figure \(9\). Phosphorylation of GS causes it to shift into its inactive conformation and inhibits glycogenesis. In addition, activated PKA also phosphorylates the protein phosphatase 1 (PP1) enzyme leading to the inactivation of the phosphatase. PP1 normally dephosphorylates GS, helping to retain the active conformation of GS. Thus, phosphorylation of PP1 by PKA helps to maintain the GS in the phosphorylated, inactive state. The inhibition of PP1, is quite complicated, as shown in Figure \(10\). PP1 contains a regulatory domain and a catalytic domain. Normally the regulatory domain of PP1 binds with glycogen, keeping the molecule close to the location where GS will be present. Thus, when GS is near its substrate it can also bind with PP1 and be dephosphorylated into its active state. This is more efficient than diffusing in the cell and trying to find the PP1 randomly. When PKA phosphorylates the regulatory domain of PP1, it dissociates from the catalytic domain, causing the catalytic domain to float away from the glycogen molecule. This makes PP1 less efficient at dephosphorylating GS because it is harder for the molecules to randomly come into contact with one another. Thus, PP1 is less active. PKA reduces this activity even further, by phosphorylating an allosteric inhibitor (I) of PP1. In the phosphorylated state, the inhibitor can bind to PP1 fully inactivating the phosphatase. Both phosphorylation events need to be reversed to regain full PP1 activity. In summary, glucagon signaling in the liver downregulates glycogenesis through the activation of PKA. PKA phosphorylates GS directly, inactivating the enzyme, and maintains it in the inactive state by also inhibiting the PP1 responsible for dephosphorylating GS. Activation of Glycogenolysis In addition to phosphorylating GS and PP1 during the inactivation of glycogenesis, PKA also phosphorylates the enzyme Phosphorylase Kinase, which is upstream of glycogen phosphorylase, the primary enzyme involved in glycogen breakdown. As its name implies, phosphorylase kinase is a protein kinase that phosphorylates the enzyme to activate it. Figure \(11\) offers a first view of the phosphorylation cascade required for glycogen phosphorylase activation. The phosphorylase kinase enzyme is a complex enzyme that is a tetramer of a tetrameric complex, αβγδ, so the full holoenzyme has an (αβγδ)4 structure. It is large with a molecular weight of around 1.3x106. As you would expect, it is highly regulated in multiple ways, including phosphorylation by PKA, ADP (an allosteric effector), divalent cations like Ca2+, and pH. The α and β are regulatory subunits that affect activity through their phosphorylation. The δ is calmodulin, a calcium-binding protein we discussed in Chapter 12.7, and its binding of calcium affects the holoenzyme activity. The γ subunit has kinase activity and has an N-terminal catalytic domain and a C-terminal calmodulin-binding domain.  This is primarily regulated by phosphorylation through the PKA pathway as shown in Figure 15.3.5. "PHK is one of the largest of the protein kinases and is composed of four types of subunit, with stoichiometry (αβγδ)4, and a total mol. wt of 1.3×106 Da. Activity is regulated by cyclic AMP-dependent protein kinase phosphorylation, autophosphorylation, allosteric effectors (e.g. ADP), metal ion concentration (Ca2+ and Mg2+), proteolysis and pH (Pickett-Gies and Walsh, 1986). The α and β subunits are regulatory and are the targets for control by phosphorylation. The δ subunit is essentially identical to calmodulin and confers Ca2+ sensitivity. The 386 amino acid γ subunit is the catalytic subunit which comprises an N-terminal kinase domain (residues 1–298) and a regulatory calmodulin-binding domain (residues 299–386)" Calcium, an allosteric regulator, may be present within cellular targets due to nerve impulse firing, muscle contraction, or through hormone signaling. The presence of calcium in the cell generally indicates that there is high energy demand on the cell at that time and that energy production is needed. Thus, calcium binding to the phosphorylase kinase is a positive effector of the enzyme and upregulates activity. Maximal activity of the enzyme is achieved through combined phosphorylation and calcium binding. Thus, phosphorylase kinase can exist in 4 different states of activity as shown in linked equilibria in Figure \(12\). This diagram by now should be quite familiar. It is yet another example of a tetrameric enzyme (where the "enzyme" composition is αβγδ) existing in two major states, an inactive T state and an active R state, with the interconversion regulated by allosteric effectors (Ca2+) and post-translationally phosphorylation. A mechanism for the phosphorylation of a Ser in a substrate target protein (i.e glycogen phosphorylase) by the catalytic domain of the γ subunit of phosphorylase kinase is shown in Figure \(13\). Figure \(13\): A mechanism for the phosphorylation of a Ser in a substrate target protein (i.e glycogen phosphorylase) by the catalytic domain of the γ subunit of phosphorylase kinase. https://www.ebi.ac.uk/thornton-srv/m-csa/entry/35/. Creative Commons Attribution 4.0 International (CC BY 4.0) License. The * in the mechanism denotes the serine of the target protein. Figure \(14\) below which shows an interactive iCn3D model of a truncated form of the rabbit phosphorylase kinase gamma subunit dimer bound to a peptide substrate complex (2PHK). Figure \(14\): Rabbit phosphorylase kinase gamma subunit dimer bound to a peptide substrate complex (2PHK). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...3wHapyHWEy7qi9 The phosphorylase kinase dimer is shown (gray and brown subunits).  A non-hydrolyzable ATP analog (adenylyl imidodiphosphate, AMPPNP) in each subunit is shown as CPK-colored sticks. The backbone of two identical peptide substrates ( blue and cyan) with the sequence RQMSFRL (similar to the target sequence in glycogen phosphorylase and an ideal peptide substrate) is shown as a backbone trace, with the central Ser which gets phosphorylated shown as CPK-colored spheres. Active site residues are shown as CPK-colored sticks and labeled in each subunit. We have been primarily discussing the regulation of glycogenolysis in the liver. However, in considering the activity of PK and its reactivity with Ca2+ ions, we should also consider the activation of glycogenolysis in skeletal muscle, as well. Of note, glycogenolysis in liver tissue and skeletal muscle has many differences. First, the G-protein coupled pathway is activated by different hormones. Liver tissue is responsive to Glucagon stimulation, as well as stimulation through the Epinephrine hormone signaling pathway. Glycogenolysis in skeletal muscle tissue, on the other hand, is only activated by the Epinephrine signaling pathway, but not by glucagon. This is because the liver is the primary organ responsible for regulating blood glucose levels. Thus, pancreatic signaling due to low blood glucose levels primarily targets glycogenolysis within the liver tissue. Both systems are responsive to Epinephrine, which is described in more detail below. Epinephrine Signaling Epinephrine is a small amino acid-derived hormone (can you guess the amino acid?? Yes it is Tyrosine!!), as shown in Figure \(15\). It is also called adrenaline, as it is secreted from the adrenal glands located just above the kidneys, during the flight or fight response. It is also secreted during heavy or sustained exercise. Epinephrine has pleiotropic responses in the body, which include the activation of glycogenolysis in the liver and skeletal muscles. Epinephrine also promotes fat breakdown in adipose tissue, which releases this energy reserve into the bloodstream for utilization by muscle tissue. It also causes the relaxation of smooth muscles in the lungs and respiratory tract enabling better oxygen absorption. Cardiac contractility is also increased to increase blood flow to skeletal muscles. This supports the generation of ATP from glucose and fatty acids for sustained muscle utilization. It also reduces blood flow to the skin and causes the contraction of smooth muscles in the skin causing goosebumps. Within the liver and skeletal muscle, the epinephrine signaling pathway overlaps with the glucagon signaling pathway seen in the liver. The epinephrine receptor is also a G-protein coupled receptor related to the glucagon receptor. However, it is specific for epinephrine and cannot bind with glucagon. It does activate the same G-protein pathway leading to Protein Kinase A activation, as shown in Figure \(16\). The body is very efficient at reusing machinery in different parts of the body, in this case, it does so under different regulatory parameters. The Glycogen Phosphorylase enzymes are also encoded by different genes within the Liver and Skeletal Muscle. These are known as isozymes. Recall, that isozymes have the same biological function, but since they are expressed from different genes, they have different enzyme kinetics and they are regulated in different and unique ways within each tissue. The liver and skeletal muscle forms of Glycogen Phosphorylase share approximately 90% sequence identity. Yet again, both isozymes can exist in two major conformations, the a-form, and the b-form. The protein adopts the a-form when it is phosphorylated at Ser 12 by phosphorylase kinase, as shown in Figure \(17\). The Glycogen Phosphorylase enzyme can also be in two different states, the relaxed, flexible state which is the active form of the enzyme, and the tense or rigid state which is inactive. When the protein is in the a-conformation, it favors the relaxed and active state of the protein. Therefore, phosphorylation of Glycogen Phosphorylase leads to an increase in the activity of the enzyme. This is depicted in the following diagram. Another way to think about these four different enzyme states is that they exist in a dynamic equilibrium, with the population of each state determined by the phosphorylation state AND the presence of allosteric inhibitors and activators which we will explore below. In Figure 17 above, the R state is on top and the T state is on the bottom. The equilibrium between just those two states is indicated by the vertical arrows. The thickness of the arrows indicates the preferred direction of the reversible reaction. In the absence of phosphorylation (left-hand vertical states), the equilibrium favors the T or inactive form. When phosphorylated (right-hand vertical states) on Ser12 by the enzyme phosphorylase kinase, the R or active form is favored. The horizontal equilibria show the phosphorylation of the Ser12 by phosphorylase kinase (top, shown reversibly but the enzyme is not acting physiologically to remove phosphate) and dephosphorylation of Ser 12 by the enzyme protein phosphatase 1 (which acts physiologically only as a phosphatase). The different isozymes of the Glycogen Phosphorylase enzyme are also regulated by different, tissue-specific allosteric effectors. Within the liver, glucose is a negative regulator of Glycogen Phosphorylase, which makes sense, as the role of this pathway in liver tissue is to promote the release of glucose into the bloodstream. The presence of free glucose in the cytoplasm of the liver would indicate either the fed-state when blood glucose levels are high, or that high levels of glycogenolysis have released substantial glucose. Within liver tissue, the presence of free glucose will cause the a-form of Glycogen Phosphorylase to shift to the Tense state, reducing the activity of the enzyme. This is shown in Figure \(18\). Skeletal muscle glycogen phosphorylase (or Myophosphorylase, as it is sometimes called), is more responsive to allosteric effectors that indicate the energy state of the cell. This makes sense, as the main purpose of glycogen breakdown in muscle tissue is to fuel the energy demand for the muscle tissue. Thus, the energy housed in glucose will be used to produce ATP within these cells. The presence of either glucose 6-phosphate or ATP within skeletal muscle indicates high levels of energy are present. Thus, glycogen breakdown will be inhibited. The presence of AMP, on the other hand, indicates a low energy state and is an activator of Glycogen Phosphorylase. This is shown in Figure \(19\) to Figure \(21\)below. Again, note that the bold red arrows point downward, showing that on the addition of glucose-6-phosphate, the T state (inactive) is favored even if glycogen phosphorylase has been phosphorylated. Again, note that the bold red arrows point downward, showing that when the ATP levels are high, the T state (inactive) is favored even if glycogen phosphorylase has been phosphorylated. Again, under conditions of a high energy state (as reflected by high ATP), there is no need to cleave glycogen to enter glycolysis. In contrast, note that the bold green arrows point upward showing that when the AMP levels are relatively high, the R state (active). Higher levels of AMP reflect a need to activate glycogen breakdown to increase ATP production. A mechanism for the phosphorolysis of Glcn+1 to Glcn and glucose-1-phosphate is shown in Figure \(22\). Note the unusual presence of a molecule of pyridoxal phosphate covalently attached through a Schiff base linkage to Lys 568 (rabbit phosphorylase). Figure \(22\): A mechanism for the phosphorolysis of Glcn+1 to Glcn and glucose-1-phosphate. https://www.ebi.ac.uk/thornton-srv/m-csa/entry/205/.  Creative Commons Attribution 4.0 International (CC BY 4.0) License PLP, which is covalently attached through a Schiff base, functions in this enzyme as a general acid and base, and not as a cofactor that facilitates covalent bond cleavage in amino acid substrates that are covalently attached to it (Chapter 6.8: Cofactors and Catalysis - A Little Help From My Friends). It is important to note that the reaction is a phosphorolysis, not a hydrolysis, which would leave free glucose, which would more readily leave the cell and hence be less available for cellular energy needs and less available for glycolysis. Figure \(23\) below which shows an interactive iCn3D model of a dimer of rabbit glycogen phosphorylase (1GDB). Figure \(23\): Dimer of rabbit glycogen phosphorylase (1GDB). (Copyright; author via source). Click the image for a popup or use this external link:https://structure.ncbi.nlm.nih.gov/i...s1LosR5TJLjru9 The subunits in the dimeric form are shown in different colors. PLP is shown in spacefill. Active site residues are shown in both subunits as CPK-colored sticks and labeled. Let's look at a monomer (from the tetramer) of 2 nonphosphorylated states of glycogen phosphorylase. Since they are both unphosphorylated, they both represent the b state. One (2GPB) has glucose bound, so it represents the inactive (T state). The other (3E3N) has AMP bound, so it represents the active (the R state). Figure \(24\) shows the conformation differences between the monomeric states Figure \(24\): Conformational differences between the monomeric unphosphorylated b state of glucose-bound GP (T state, inactive) and AMP-bound GP (R state, active) Now let's look at the conformations of two different GP activated by different means. In one, the nonphosphorylated form of GP (the b state, and inactive T state, ) binds AMP, an allosteric activator (pdb 8GPA), and converts to the active, b state (unphosphorylated R state). Let's compare its active conformation to a form of GP activated by phosphorylation (the phosphorylated a state, and active R state) to which just SO42- is also bound. In the 1st case, GP-b T state is driven to the active Gp-b R state by binding of the allosteric activator AMP. In the second case, GP-a is already in the R active state since it is phosphorylated.  Figure \(25\) compares their conformations. Figure \(25\): Comparison of the conformations of two active forms of GP - phosphorylase b bound to the allosteric activator AMP (R state) and phosphorylase a activated by phosphorylation of Ser 14. The cyan monomer is glycogen phosphorylase b which is not phosphorylated but driven into the R state on the binding of AMP (note that two are bound at the periphery). The dark blue monomer is glycogen phosphorylase a which is phosphorylated at Ser14 (a different number in this crystal file which is shown in spacefill and labeled SEP-14) which is also in the R state. It also has two SO42- bound that help stabilize the state. Look carefully! the conformations are very similar to each other, in contrast to the different conformations for the T and R states shown in Figure 24. Figure \(26\) below which shows an interactive iCn3D model of unphosphorylated (b state) rabbit glycogen phosphorylase with bound glucose (inactive T state, 2GPB). Figure \(26\): Unphosphorylated (b state) rabbit glycogen phosphorylase with bound glucose (inactive T state, 2GPB). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...eXQciAPczmQpLA Only 1 monomer of the tetramer is shown. PLP and glucose are shown in spacefill, CPK colors, and labeled. Key active site residues are shown as colored sticks and labeled. Figure \(27\) below which shows an interactive iCn3D model of unphosphorylated (b state) rabbit glycogen phosphorylase with bound AMP (active R state, 3E3N). Figure \(27\): Unphosphorylated (b state) rabbit glycogen phosphorylase with bound AMP (active R state, 3E3N). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...WDZJc1CcpFqEG7 Note the different locations for the binding site of the allosteric activator AMP compared to the inhibitor glucose in the previous model. McArdle's Disease McArdle's Disease, also referred to as myophosphorylase deficiency or type V glycogen storage disease, is a recessive inherited disorder characterized by an inability to metabolize glycogen due to the absence of a functional myophosphorylase (PYGM). In Figure \(28\) shown below, the normal functional pathway is shown in blue, on the left, while the mutant pathway is shown on the right in red. Patients with this disease lack sufficient glucose-1-phosphate (G1P) monomers needed for glycolysis and the hexosamine biosynthetic pathway (HBP). This results in lower ATP and, consequently, lower muscle contraction, as well as in lower post-translational modifications by O-GlcNAcylation in comparison to normal conditions. This is especially pronounced during extended or heavy workouts, where people with McArdle’s Disease will sustain painful cramping of their muscle tissue during workouts, can have dark red/brown urine, and can easily tire during activity. Some patients also note a second-wind phenomena occur during workouts as the body shifts from carbohydrates to lipids as a primary energy source. The dark red/brown color in the urine happens if muscle tissue is damaged during the workout. The damaged muscle releases the protein myoglobin into the bloodstream. This is filtered out by the kidneys and excreted in the urine, causing the color change. The severe uncontrolled disease can cause life-threatening kidney problems. Figure \(29\) presents an overview of glucose metabolism in skeletal muscle. Both glucose-1-phosphate (G1P) released from the intracellular glycogen stores by glycogen phosphorylase (GP), as well as the glucose introduced into the cell through glucose transporters (GLUT) are converted to glucose-6-phosphate (G6P) by phosphoglucomutase (PGM) and hexokinase (HK), respectively. The G6P can be directed to different destinations. One of them is the pentose phosphate pathway for the formation of nucleic acid building blocks (ribose and deoxyribose). Another is in the formation of ATP. Here G6P enters the metabolic pathway of glycolysis. The glycolytic reactions culminate in the production of pyruvate and adenosine triphosphate (ATP). Pyruvate can be fermented to lactate by the catalysis of the lactate dehydrogenase (LDH), as happens during anaerobic muscle exercise. On the other hand, pyruvate can be used to obtain ATP through full oxidation in the Kreb Cycle. In total, oxidative phosphorylation produces between 30-36 molecules of ATP (depending on the organism and tissue), 6 molecules of carbon dioxide (CO2), and 6 molecules of water (H2O) from 1 glucose molecule. Glycolysis alone only produces two net ATP molecules per glucose. Glucose, in addition to being the main fuel of the cell’s energy metabolism, is also used by the cellular machinery as a vitally important substrate for the production of key intermediaries of the hexosamine biosynthetic pathway (HBP) forming O-GlcNAc, β-linked N-acetylglucosamine. And finally, in times of plenty, glucose will be utilized by glycogen synthase (GS) to make glycogen.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/15%3A_Glucose_Glycogen_and_Their_Metabolic_Regulation/15.03%3A_15.3_Glycogenolyis_and_its_Regulation_by_Glucagon_a.txt

Search Fundamentals of Biochemistry There are three major enzymatic control points within the glycolytic pathway. These include hexokinase, phosphofructokinase, and pyruvate kinase reactions. Key drivers for regulating the pathway are energy demand within the cell as determined by local indicators such as ATP and AMP, as well as energy demand within the organism as a whole, which can be influenced by hormone signaling pathways. We will also see that the regulation of the pathway can vary depending on cell type and cellular needs. Hexokinase Regulation One of the primary mechanisms that control the regulation of the hexokinase step in glycolysis is the presence of different hexokinase enzymes in different cellular types. Essentially, these are proteins that are encoded by different genes but perform the same function within the cell. They are known as isozymes. Isozymes can have different enzyme kinetics, different expression patterns in different tissues, different post-translational modifications, and bind with different allosteric effectors. This affords the body to have differential control over the same processes in different locations within the body. Within vertebrates, four important hexokinase isozymes vary in subcellular locations and kinetics parameters. This allows the differential phosphorylation of hexoses depending on local conditions, and physiological function. They are designated hexokinases I, II, III, and IV. All of the Hexokinases can use multiple hexoses as substrates, in addition to glucose. These include mannose, fructose, and 2-deoxyglucose. Hexokinase IV is also often referred to as Glucokinase and is specific to the Liver and Pancreas. Recall that hexokinase enzymes mediate the first step in the glycolytic pathway with the formation of glucose 6-phosphate, as shown in Figure \(1\) below.  They also require ATP as a cofactor in the process. Recall that glucose-6-phosphate (G6P) has several potential fates within the body, as shown in Figure \(2\). It can be used as an energy source through the pathways of glycolysis and aerobic respiration. Short bursts of anaerobic respiration can also be sustained in animals that convert pyruvate into lactate. G6P can also be dephosphorylated in the liver and released back into the bloodstream to maintain homeostatic balance. The pancreas uses G6P as a sensor to determine when to secrete insulin and glucagon. The G6P can also serve as a building block for anabolic processes. It can be converted to ribose through the Pentose Phosphate Pathway where it will be used in the construction of nucleotide monomers. It can also be used for the formation of hexosamines used in proteoglycan and glycoprotein formation. Now let’s take a look at the different isozymes of hexokinase in a little more detail. Hexokinase I (HKI) is found widely distributed throughout the body and is the main form expressed in brain tissue and red blood cells, as shown in Figure \(3\). In brain cells, this protein is localized to the mitochondria. This colocalization aids in the efficient coupling of glycolysis and the Krebs' cycle and oxidative phosphorylation pathways inside the mitochondria. It also ties the activity of HKI with oxidative phosphorylation and energy load, as HKI preferentially uses mitochondrially-derived ATP in its reaction mechanism. HK association with the mitochondria also has a cellular protective effect, reducing the potential for programmed cell death or apoptosis to occur. Red Blood Cells (RBCs), on the other hand, are highly differentiated cells with a very short lifespan. They are replaced in humans approximately every two weeks. RBCs are enucleated and do not have mitochondria, and thus, only generate ATP through the process of glycolysis. the HKI protein is free floating in the cytoplasm in this system. HKI has a low Km, meaning that it has a high affinity for glucose and is active at low substrate concentrations. It is also inhibited by the product Glucose-6-Phosphate (G6P) in the process of negative feedback inhibition. Essentially, you do not want to waste time and energy making more than you need. Low to moderate levels of free inorganic phosphate can overcome this negative feedback inhibition. HKI and HKII are expressed in Skeletal Muscle, Heart Muscle, and insulin-sensitive tissues. While it is thought that HKI is providing a predominantly catabolic role for the use of G6P in energy production, HKII may play a more pertinent role in anabolic processes, providing G6P for conversion to G1P and subsequent utilization in Glycogenesis. Both HKI and HKII are localized to the outer membrane of the mitochondria. However, while 95% of HKI is associated with mitochondria, only about 70% of HKII is associated, with the remaining HKII fractionating with the cytosolic proteins. This could help to explain the heightened role of HKII in anabolic glycogenesis processes within skeletal muscle, and why it is not found in brain tissue. HKII is also often overexpressed in tumor cells, where it is associated with higher mortality rates. It has also been linked with the processes of metastasis and with the development of drug resistance. Similar to HKI, HKII also has a low Km and is inhibited by G6P, although this inhibition is not released by the presence of inorganic phosphate. Not a lot is known about the functions of HKIII. It may be an inactive gene duplication or remnant. Under basal conditions, it is not expressed to appreciable levels in any major tissues, and studies on its biological activity it is inhibited by glucose at physiological concentrations. However, some studies suggest that it may be expressed during cellular stress responses, such as hypoxia, although its function in these types of responses is not currently understood. Hexokinase IV or Glucokinase is specifically expressed within the liver and pancreas. HKIV is cytoplasmic and not tethered to the mitochondria. Activity within the pancreas serves as a sensor for the release of insulin, and in the liver for the production of G6P that will fuel glycogen production. HKIV has a higher Km than HKI and HKII, thus it does not work efficiently at low concentrations of glucose. However, it is NOT inhibited by the product, G6P. Thus, it will continue to make G6P, even when levels are high. This helps to explain the high levels of glycogen that are stored within liver tissue, but not elsewhere in the body. This also ensures that the sensor system in the pancreas will accurately read blood glucose levels. The four isozymes of HK share high homology with one another and appear to have arisen from gene duplication events, as shown in Figure \(4\). The left-hand panel of Figure 15.5.4 shows the linear protein domains of the different HK isozymes. Both HKI and HKII contain an N-terminal domain that localizes the protein to the mitochondrial membrane. HKI, II, and III all contain two repeating catalytic domains in the N- and C-terminals. However, mutations in the N-terminal domain in HKI and HKIII render them inactive. Both catalytic domains in HKII retain activity. HKIV (Glucokinase) is the most truncated isozyme, only containing the C-terminal catalytic domain. The lower right diagram shows the ribbon diagram of HKIV. Upstream promoter regions of HKIV (not shown in this diagram) also differ, allowing for controlled expression in the liver and pancreas. Feedback inhibition in HKI and HKII occurs through the N-terminal catalytic domain. The upper diagram on the right shows an HKI dimer complex with an ATP analog, glucose, G6P, and Mg2+ ion. HKI dimerizes when concentrations of the inhibitor, G6P, are high enough. Dimerization reduces the biological activity of the enzyme in brain tissue. Dimerization of HKI can be reversed in the presence of low levels of inorganic phosphate. HKIV is also a good model for understanding enzyme conformation change during the reaction. HKs change shape by induced fit upon substrate binding. HKIV has a large induced fit motion that closes over the substrates ATP and xylose, as shown in Figure \(5\).  The binding sites are shown in blue, substrates in black, and the Mg2+ cofactor in yellow (PDBs:2E2N,2E2Q). In summary, the isozyme expression patterns of HKs differentially regulate the fate of glucose within those tissues. Within brain tissue and red blood cells where only HKI is present, glucose is predominantly used in the glycolytic pathway for energy production. In muscle tissue, the presence of HKII allows for increased use of glucose for the formation of glycogen. HKIV expression in the pancreas and liver allows for the homeostatic regulation of blood glucose levels and stockpiling of glucose in the form of glycogen. Phosphofructokinase-1 Regulation Recall that phosphofructokinase-1 (PFK1) mediates the third step in the glycolytic pathway with the conversion of fructose 6-phosphate to fructose 1,6-bisphosphate, as shown in Figure \(6\).  The PFK1 reaction is the first irreversible reaction of glycolysis. It also represents the committed step within the pathway. The phosphorylation of fructose-6-phosphate (F6P) to fructose-1,6-bisphosphate (F1,6BP) commits the F1,6BP to continue through the glycolytic pathway. It cannot be utilized for any other purpose at that point. F6P, on the other hand, could be converted back into glucose-6-phosphate and used for many different purposes (ie glycogen synthesis, nucleotide synthesis, or hexosamine synthesis). Because of the committed nature of this step, PFK1 is one of the most important control points in the glycolytic pathway. The PFK1 enzyme is composed of a tetramer that can contain different combinations of three types of subunits: muscle (M), liver (L), and platelet (P). The composition of the PFK1 tetramer differs according to the tissue type it is present in. For example, mature muscle expresses only the M isozyme, therefore, the muscle PFK1 is composed solely of homotetramers of M4. The liver and kidneys express predominantly the L isoform. In erythrocytes, both M and L subunits randomly tetramerize to form M4, L4, and the three hybrid forms of the enzyme (ML3, M2L2, M3L). As a result, the kinetic and regulatory properties of the various isoenzymes pools are dependent on subunit composition. Tissue-specific changes in PFK1 activity and isoenzymes content contribute significantly to the diversities of glycolytic and gluconeogenic rates which have been observed for different tissues PFK1 is an allosteric enzyme and has a structure similar to that of hemoglobin in so far as it is a dimer of a dimer, as shown in Figure \(7\).  One half of each dimer contains the ATP binding site, whereas the other half the substrate (fructose-6-phosphate or (F6P)) binding site, as well as a separate allosteric binding site, that can bind with ADP or AMP. All isoforms of PFK1 are activated by the allosteric binding of ADP or AMP. This indicates a low energy state within the cell and the need for glycolysis and energy generation. Allosteric inhibitors include high levels of ATP and Citrate. Note that ATP is a substrate of this enzyme and has the normal substrate binding site. When there is enough ATP present that it can also bind allosterically to the enzyme, it will act as an inhibitor. Citrate, the first molecule in the Kreb's Cycle (Citric Acid Cycle), can also act as an allosteric inhibitor of PFK-1. High levels of citrate that no more pyruvate is needed to generate ATP through oxidative phosphorylation. We are also going to see that Fructose 2,6-bisphosphate is predominantly a regulator of PFK1 in Liver Cells, where it serves as an activator. Before we discuss the formation and use of Fructose 2,6-bisphosphate and its role in the regulation of PFK1, let’s review opposing glucose utilization/production pathways within the liver, as shown in Figure \(8\). Two opposing pathways within the liver are glycolysis (the breakdown of glucose) and gluconeogenesis (the formation of glucose). It would be unproductive to have both of these pathways operating at the same time. Thus, if one pathway is needed, then the other one needs to be turned off. We previously saw this same type of control for glycogenesis (the formation of glycogen) and glycogenolysis (the breakdown of glycogen). When one of these pathways is upregulated, the other needs to be downregulated. In the resting state of the liver, when blood glucose levels are in the homeostatic range, gluconeogenesis will be shut off, as it is very expensive to make glucose and the liver will only invest in making glucose if blood glucose levels fall critically low (ie extreme exercise or long term fasting). In the resting state, the liver will use the glycolytic pathways to supply normal levels of ATP energy for the maintenance of housekeeping processes. Glycogenesis and glycogenolysis will be in balance or equilibrium as needed to augment the supply of glucose entering the system from the bloodstream. If glucose in the blood drops below homeostatic levels, the pancreas will release glucagon and begin this hormone-signaling pathway that causes the liver to release glucose into the bloodstream. Thus, glucagon signaling leads to the downregulation of glycolysis and glycogenesis, so it can shunt glucose pools to the bloodstream. It also leads to an increase in glycogenolysis or the breakdown of glycogen. During this time, liver cells are predominantly generating ATP from lipids, rather than carbohydrates. Thus, glycolysis can be inhibited to promote the release of glucose into the bloodstream. If cellular demand for glucose is high, liver cells will also turn on the gluconeogenesis pathway and make glucose new from non-carbohydrate precursors (which it then exports into the bloodstream for delivery to tissues). This is an energy-intensive pathway. The major site of gluconeogenesis is the liver, with a small amount also taking place in the kidney. Little gluconeogenesis takes place in the brain, skeletal muscle, or heart muscle. It just does not make energetic sense to do that! It costs more energy for cells to make glucose than they can get from breaking it down in oxidative phosphorylation. This cost can be dealt with by doing it in the liver and then releasing it into the bloodstream to fuel activities in the brain, heart, and skeletal muscles. To do this, glucagon stimulates that lovely signaling pathway that you are all now familiar with (reviewed in Figure \(9\))! Within the liver, it activates the G-protein coupled receptor which in turn activates the downstream G-protein. Adenylate Cyclase is activated and produces the second messenger, cAMP. cAMP binds with the CREB protein and activates the transcription of proteins involved in gluconeogenesis. The cAMP also binds with Protein Kinase A and upregulates the activity of glycogen phosphorylase resulting in the breakdown of glycogen. It also down-regulates the activity of glycogen synthase to inhibit glycogenesis. In this section, we will also see how PKA activation will down-regulate the activity of the glycolytic pathway as well. Regulation of the glycolytic pathway in response to this signaling occurs through the regulation of the PFK-2/FBPase-2 Enzyme. The activity of this enzyme is controlled through the PKA signaling cascade. This enzyme is responsible for phosphorylating fructose 6-phosphate to the fructose 2,6-bisphosphate form. Note that this bisphosphate form of fructose is DIFFERENT than the bisphosphate form utilized in the glycolytic pathway. The glycolytic pathway requires fructose 1,6-bisphosphate that is formed from the PFK1 enzyme (or Step 3 of the glycolytic pathway). The PFK-2/FBPase-2 is a separate enzyme altogether and not involved directly in the glycolytic pathway. However, we noted previously that fructose 2,6-bisphosphate can serve as an allosteric activator of the PFK1 enzyme. The PFK-2/FBPase-2 Enzyme is a dual-purpose enzyme, as shown in Figure \(10\). Half of the protein has kinase activity and can phosphorylate fructose-6-phosphate to fructose-2,6-bisphosphate. The other half of the enzyme contains a phosphatase that can cleave off the phosphate group from the 2-position and restore fructose-6-phosphate. Note that both activities are not functional at the same time! The enzyme activity is determined by the phosphorylation state. When the protein is dephosphorylated, the PFK-2 enzyme is active, leading to the production of fructose-2,6-bisphosphate. This molecule can then bind with PFK-1 in the glycolytic pathway and increase its activity. If the protein is phosphorylated by PKA during glucagon signaling, the FBPase is activated and the kinase activity is inhibited. This leads to the dephosphorylation of fructose at the 2-position and the release of fructose-6-phosphate. As shown in Figure 15.5.10, the PFK-2/FBPase-2 Enzyme is responsible for phosphorylating fructose 6-phosphate to the fructose 2,6 bisphosphate form. Note again that this bisphosphate form of fructose is DIFFERENT than the bisphosphate form utilized in the glycolytic pathway. The glycolytic pathway requires fructose-1,6-bisphosphate that is formed from the PFK1 enzyme (or Step 3 of the glycolytic pathway). If there is a lot of fructose-6-phosphate around (ie you just drank high fructose corn syrup in your sugary energy drink) it can support both the formation of fructose 1,6-bisphosphate by PFK1 and the production of fructose 2,6-bisphosphate by PFK-2. Fructose 2,6-bisphosphate will then bind with PFK1 and increase its activity converting fructose 6-phosphate into fructose-1,6-bisphosphate, as shown in Figure \(11\). However, during glucagon signaling, you need to shut down this fast-track upregulation of PFK1 by fructose-2,6-bisphosphate and turn down the glycolytic pathway. To do this, Protein Kinase A will phosphorylate the PFK-2/FBPase-2 enzyme and alter its activity. The kinase activity is inhibited and the phosphorylase activity is turned on. Figure \(12\) provides a summary of this pathway control. In the presence of glucagon, PKA will phosphorylate the PFK-2/FBPase-2 enzyme causing the kinase activity to be switched off and the phosphatase activity to be switched on. Dephosphorylation of fructose-2,6-bisphosphate recovers fructose-6-phosphate (F6P). F6P can then go through the reverse isomerase reaction and recover glucose-6-phosphate (G6P). G6P will then be transported to the rER where it will be dephosphorylated and then free glucose can be released back into the blood. The opposite holds for insulin signaling. High insulin concentrations result in the activation of protein phosphatase and the dephosphorylation of the PFK-2/FBPase-2 enzyme. In the dephosphorylated state, PFK-2 activity is high and the FBPase-2 activity is low, which will stimulate PFK1 and the glycolytic pathway. Pyruvate Kinase Recall that pyruvate kinase mediates the final reaction during glycolysis resulting in the production of pyruvate and ATP as shown in Figure \(13\). Similar to PFK1 this enzyme is also a key regulatory component within the pathway. The pyruvate kinase enzyme exists as a tetramer, that is built from the combination of different isozymes expressed in different tissues. There are three major isozymes of pyruvate kinase, the L form that is predominantly found in the liver, the R form that is predominantly found in erythrocytes, and the M1 form in muscle and brain, and the M2 form that is expressed in fetal tissue and at some level in most adult tissues. The L and R forms are splice variants that arise from the same gene locus, and the M1 and M2 forms are also splice variants that arise from the same gene locus. We will focus on some of the general regulatory mechanisms common to most of the isozymes of pyruvate kinase, starting with the activator, fructose 1,6-bisphosphate (FBP). Because FBP is an earlier product within the same metabolic cascade, the activation of pyruvate kinase enzymes by FBP is known as feedforward stimulation. All of the isozymes, except for the M1 form are stimulated by the binding of FBP to the enzyme. Similarly, all of the pyruvate kinase isozymes are inhibited by the product of the reaction, ATP (or high energy load), and high levels of alanine. Alanine can be converted to pyruvate in one enzymatic step. Thus, pyruvate serves as a metabolic intermediate in the formation of alanine. If high levels of alanine are present, this indicates that there is a high energy load within the cell (ie that the cell is full of building blocks to make new macromolecules and is not in the need of more energy). Thus, high levels of alanine serve as a negative regulator of the pyruvate kinase family of enzymes. The liver isozyme of pyruvate kinase is also regulated through protein phosphorylation, as shown in Figure \(14\). Similar to the PFK-2 activity of the PFK-2/FBPase-2 enzyme, the liver isozyme of Pyruvate Kinase is also downregulated during glucagon signaling. Protein kinase A phosphorylates Pyruvate Kinase inhibiting its activity and preventing the conversion of phosphoenolpyruvate to pyruvate. Dual regulation of the glycolytic pathway during glucagon signaling helps to ensure that glucose resources will be diverted away from cellular use by the liver and released into the blood stream to restore homeostatic blood glucose levels. Fructose Regulatory Bypass Other sugars from the diet can also enter into the glycolytic pathway, as shown in Figure \(15\). Galactose is converted in a four-step process to Glucose-6-phosphate and mannose can be converted to fructose-6-phosphate. Within most of the body’s tissues, fructose can also be converted into fructose-6-phosphate by hexokinase. However, in the liver and kidneys, there is an alternative route that fructose from the diet can take to enter into the glycolytic pathway. This pathway is concerning because it bypasses two of the major regulatory steps of the glycolytic pathway, the hexokinase step and the PFK1 step. Within the liver and kidneys, fructose can also be converted into fructose-1-phosphate by the enzyme fructokinase. The other isozyme of Aldolase, Aldolase B, can cleave the fructose-1-phosphate into 2 three carbon units, dihydroxyacetone phosphate, and glyceraldehyde. Dihydroxyacetone phosphate can be converted to glyceraldehyde 3-phosphate by Triose Isomerase and then continue into the glycolytic cascade. Glyceraldehyde can be phosphorylated to Glyceraldehyde 3-phosphate by Triokinase. This is an unregulated system that can flood the Kreb Cycle with high levels of pyruvate if high levels of fructose enter the cell (i.e. from high fructose corn syrup, sucrose, and other sweeteners common to the Westernized diet). The excess pyruvate can then be shunted into fatty acid biosynthesis for long-term storage in the form of triglycerides. If the pathway is overutilized by consuming too much sucrose and high fructose corn syrup, this can lead to the development of Hypertriglyceridemia (or the heightened increase of body fat). With this, we will end our discussions of glycolysis. In the next section, we will look at the complementary and opposite pathway, gluconeogenesis. 15.05: Regulation of Gluconeogenesis Search Fundamentals of Biochemistry Within the regulation of the gluconeogenic pathway, three of the major enzymatic steps are regulated. The first two are the pyruvate carboxykinase enzyme and the phosphoenolpyruvate carboxykinase (PEPCK). Recall that these two enzymes are required to convert pyruvate back into phosphoenolpyruvate via an oxaloacetate intermediate as shown in Figure \(1\). The third enzyme regulated in this pathway is Fructose 1,6-Bisphosphatase which converts fructose 1,6-bisphosphate into fructose 6-phosphate. We will explore the regulation of these three enzymes in more detail. Pyruvate Carboxykinase Pyruvatecarboxykinase is one of the primary regulation points. It is primarily regulated by two allosteric effectors, Acetyl-CoA and ADP, as shown in  Figure \(2\). When pyruvate enters into the Kreb Cycle, it is first converted to Acetyl-CoA. If abundant pyruvate is present, an ample supply of acetyl-CoA will also be available, indicating a high energy load for the cell. Acetyl-CoA, can bind with pyruvate carboxylase and act as an activator of the protein, stimulating the production of oxaloacetate. ADP, on the other hand, is a low-energy indicator and an inhibitor of the enzyme. In the next section, we will discover how oxaloacetate moves into the cytoplasm. Phosphoenolpyruvate Carboxykinase Cytoplasmic PEPCK is largely regulated at the transcriptional level. Increases in gene expression are seen in response to elevated cAMP levels, increased glucocorticoids, and increased thyroid hormone levels, as shown in Figure \(3\). The activated CREB transcription factor plays a role in this response. Alternatively, decreased gene expression is caused by insulin signaling. ADP also acts as an allosteric effector of the protein, causing it to have lower activity. This indicates that when energy is low, the cell cannot afford to use its reserves to remake glucose and inhibits the pathway. Fructose 1,6-Bisphosphatase Fructose 1,6-bisphosphatase is both competitively and allosterically regulated, as shown in Figure \(4\). Fructose 2,6-bisphosphate serves as a competitive inhibitor of the enzyme reducing the overall activity of the enzyme for fructose 1,6-bisphosphate. Competitive inhibitors bind within the active site and compete for binding with the regular substrate. Thus, they lower the overall Km of the reaction and make the enzyme less effective at lower substrate concentrations. However, the Vmax of the enzyme is not affected during the process. In addition to competitive inhibition, low energy load (AMP and ADP) also inhibits the enzyme. ADP and AMP will bind allosterically with the enzyme and inhibit its activity.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/15%3A_Glucose_Glycogen_and_Their_Metabolic_Regulation/15.04%3A_Regulation_of_Glycolysis.txt

The citric acid cycle – also known as the TCA cycle or the Krebs cycle – is a series of chemical reactions to release stored energy through the oxidation of acetyl-CoA derived from carbohydrates, fats, and proteins. 16: The Citric Acid Cycle Search Fundamentals of Biochemistry Introduction Let's do a short review of the metabolic processes for the extraction of energy from the oxidation of glucose (i.e. glycolysis). Glycolysis is a universal pathway used to anaerobically extract energy from glucose, a six-carbon. In this linear pathway, 2- 3 carbon molecules of pyruvate are formed as glucose is cleaved and converted to two molecules of glyceraldehyde-3-phosphate, form through an oxidation reaction using the oxidizing agent NAD+. As glycolysis continues, NADH builds up. Using lactate dehydrogenase, pyruvate, the end product of glycolysis, can be converted to lactate, regenerating NAD+ so the pathway can continue. The reactions are illustrated in Figure $1$. Figure $1$: Anaerobic production of pyruvate and lactate A careful glance at the structure of the 3-carbon pyruvate molecule shows that much more energy could be extracted from it, presumably through oxidative decarboxylation reactions, converting the carbons to 3 CO2 molecules. A problem arises immediately when examining pyruvate. It is an α-ketoacid and there is no easy route to decarboxylate it as an electron "sink" is not available to receive the electrons and in the process stabilize the transition state and intermediate in the reaction. This stands in contrast to the decarboxylation of β-keto acids, which have a built-in electron "sink", an electronegative carbonyl carbon, to receive the electrons. This is illustrated in Figure $2$. Figure $2$: Comparison of the decarboxylation of α and β keto acids To begin the process of complete oxidation of the remnants of glucose, pyruvate enters the mitochondria and starts the process of oxidative decarboxylations by interacting with the pyruvate dehydrogenase complex (PHC). This catalyzes a complicated reaction to attach an electron "sink" beta to the carboxylate, which is subsequently released as CO2. The end products of the PHC oxidative decarboxylation reaction are the two-carbon acetyl-CoAs, NADH, and CO2. The acetyl-CoA then enters a cyclic, non-linear pathway called the citric acid cycle, tricarboxylic acid (TCA) cycle, or Kreb's cycle, named after Hans Krebs who discovered it. We'll talk about that in section 16.2. A glance reveals that we have taken glucose a small fraction along the way of oxidizing every carbon in it to CO2 and H2O. Complete oxidation happens under aerobic conditions when the glycolytic pathway is followed by the Kreb's cycle. Pyruvate formed in glycolysis enters the mitochondrial matrix where it gets oxidatively decarboxylated while reacting with a small thiol, Coenzyme A (CoASH) to form a 2C "activated acetate" acetyl group connected through a thioester link to CoASH, forming acetyl-CoA. The third carbon from pyruvate is released as CO2. The reaction is catalyzed by the enzyme pyruvate dehydrogenase complex (PDC). Pyruvate Dehydrogenase Mechanism This enzyme complex is enormous. The E. Coli complex has a molecular weight of almost 4 million with at least 16 chains each of three different enzymes catalyzing part of the reaction. The components are pyruvate dehydrogenase (E1), dihydrolipoamide dehydrogenase (E2), and dihydrolipoamide dehydrogenase (E3). The molecular weight of the bovine complex is almost 8 million, and it has 22 E1, 60 E2, and 6 E3 subunits. Nature often uses the same solution for identical problems. For example, many proteases have an active site nucleophilic serine, which works with the assistance of histidine and aspartate to cleave peptide bonds. There are three α-ketoacid dehydrogenase complexes in many organisms. Each has a common E3 but specific E1 and E2 enzymes. Figure $3$ shows an image of the structure so you can get an overview before we dive into the activity of each of the substrates. Figure $3$: View of pyruvate dehydrogenase. https://electron.med.ubc.ca/2018/07/...dehydrogenase/ The E3 subunit is not readily seen in the image below. Why has nature produced such a monstrous enzyme complex to simply catalyze the oxidative decarboxylation of a small three-carbon molecule? We will explore that at the end of this section. The complex also employs collectively 5 substrates/cofactors derived from vitamins. • Thiamine in the form of thiamine pyrophosphate (TPP), which is covalently attached to E1 • lipoic acid, in the form of lipoamide, which is covalently attached to a lysine side chain in E2 • riboflavin the in the form of flavin adenine dinucleotide (FAD/FADH2), which is bound very tightly (and not released) to E3 • pantothenic acid, incorporated into the structure of CoASH/Acetyl-CoA, a substrate/product pair for the reaction • niacin, nicotinic acid, in the form of NAD+/NADH, a substrate/product pair for the reaction The structures for the five are shown in Figure $4$, along with some additional descriptions that summarize some of the chemistry of these molecules. Figure $4$: Structure of the cofactors in pyruvate dehydrogenase Figure $5$ shows a schematic of the overall reaction. Figure $5$: Overall reactions catalyzed by pyruvate dehydrogenase The net reaction is $\ce{pyruvate + CoASH + NAD^{+} -> Acetyl-CoA + CO2 + NADH + H^{+}} \nonumber$ Part 1: Oxidative Decarboxylation - pyruvate dehydrogenase (E1p) So let it begin. We need to get rid of one carbon as CO2 and transfer the other two carbons of pyruvate to CoASH to form acetyl-CoA, the thioester of CoASH. Thioesters are "high energy" with respect to their hydrolysis products as the thioester is destabilized compared to a normal carboxylic acid ester.  (Remember, there is no such thing as a "high energy" bond). Since the sulfur atom is larger than the O in the C-S and C-O bond in their respective esters, the thioester as a reactant can not be stabilized well as the C-S single bond length is longer, as shown in Table $1$ below. bond length (Angstroms) C-O 1.43 C=O 1.21 C-S 1.82 C=S 1.56 Table $1$: Bond lengths of carbon-oxygen and carbon-sulfur single and double bonds This minimizes resonance stabilization compared to the carboxylic acid ester, as shown in the figure below. The products of hydrolysis of both a carboxylic and thiol ester are of comparable energy. Hence only the thioester is relatively destabilized compared to its hydrolysis product, with the ΔG0 hydrolysis = -7.5 kcal/mol (-31 kJ/mol), the same as for the hydrolysis of a phosphoanhydride bond of ATP. Additionally, a resonance structure shows a positive charge on the carbonyl C and a negative on the oxygen allowing the carbonyl carbon to be more electrophilic. Another more sophisticated reason for the relative destabilization of the thiol ester is that the overlap between the carbonyl C p orbital is the larger S p orbital is less, hindering the delocalization of electrons needed to stabilize the thiol ester. Figure $6$ illustrates these points. Figure $6$: Comparison of resonance stabilization of carboxylic esters and thioesters Now we can explore the mechanism of CO2 release and acetyl-CoA production by E1. The carbon atom directly between the N and S in the ring has a reduced pKa, so it can be deprotonated to form a carbanion. The negative charge can't be stabilized by resonance but it is adjacent to the positively-charge N, which stabilizes it. This zwitterion is called an ylide, which is a net neutral species with a positive charge (usually on a N, P, or S) and a negative charge (usually on a C) on adjacent atoms. The carbanion on the ylid attacks the electrophilic C=O of pyruvate, forming a TPP intermediate with a wonderful electron sink (N+) beta to the carboxyl carbonyl C. This is the essence of the entire reaction as this enables the decarboxylation event. The rest of the reactions catalyzed by E2 and E3 allow the release of the other 2 Cs of pyruvate as acetyl-CoA (E2) and the return of the enzyme to its original state (E2 and E3). Figure $7$ shows the reaction mechanism of E1. Figure $7$: Reaction mechanism of E1 of pyruvate dehydrogenase Note that the carbonyl C in pyruvate has two single bonds to two other carbon atoms, while in the final covalently attached form it has one bond to carbon and one to sulfur. Sulfur is under oxygen in the periodic table so by analogy, the replacement of one C-C bond with a C-S bond is an oxidation reaction, which requires an oxidizing agent. The covalently attached ring of the lipoamide with an S-S bond similar to that of a disulfide bond, an oxidized form of sulfur, is the oxidizing agent. On the formation of acetyl-lipoamide, the S-S bond is cleaved and a thioester is formed. Other sulfur is a free reduced thiol. There are 22 E1 subunits in the bovine PDC. Here is an iCn3D model of one human pyruvate dehydrogenase E1 component complex that has TPnP (TDP acetyl phosphonate, a TPP analog, covalently attached (PDB ID: 6CFO). One E1 subunit is an α2β2 heterodimer. The two alpha chains are shown in cyan while the beta chains are in dark blue. Orient the model to view along the C2 rotational symmetry axes shown. Figure $8$ shows an interactive iCn3D model of the human pyruvate dehydrogenase E1 covalently bound to TDP acetyl phosphonate (TpnP), a TPP analog (6CFO) Figure $8$: Human pyruvate dehydrogenase E1 covalently bound to TDP acetyl phosphonate (TpnP), a TPP analog (6CFO). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...7FJ2GE47tvS6g7 A heterotetramer containing 2 α (cyan) and 2 β chains (dark blue) is shown. Figure $9$ shows an interactive iCn3D model of the TPP analog covalently bound to E1 of pyruvate dehydrogenase (6CFO) Figure $9$: TPP analog covalently bound to E1 of pyruvate dehydrogenase (6CFO) . (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...SNf5sC7SqmQkc8 The dotted lines show the interactions between the TPP analog, color-coded as shown in the legend below. Parts 2 and 3: Formation of Acetyl-CoA (E2) and Regeneration of the Active Complex. The next part of the reaction produces acetyl-CoA (E2), but after that, the enzyme is "dead" as it no longer has an oxidized form of lipoamide to serve as an oxidizing agent (which gets reduced) in another round of catalysis. To regenerate enzyme activity, the reduced lipoamide, after the release of the attached acetyl group, must be reoxidized by another oxidizing agent. That oxidizing agent is FAD, which is covalently attached to E3, and is converted to FADH2. It must be reoxidized back to FAD to restore activity to the enzyme complex. The final oxidizing agent used for that is solution-phase NAD+, which is released by the enzyme as a product. So it's a bit complicated. Three oxidizing agents are used in the PDH, two of which are covalently attached to the enzyme (oxidized lipoamide on E2 and oxidized FAD on E3). Figure $10$ shows the transacetylation reaction and formation of reduced lipoamide Figure $10$: Transacetylation reaction and formation of reduced lipoamide by pyruvate dehydrogenase E2 The reaction of E3 follows to restore the fully catalytic enzyme, as shown in Figure $11$. Figure $11$: Regeneration of oxidized lipoamide by pyruvate dehydrogenase E3. Let's look in greater detail at the structures of both E2 and E3. E2: dihydrolipoyl acetyltransferase - In the mammalian complex, 60 E2 subunits arranges into a pentagonal dodecahedron. Most gram-negative bacteria E2 subunits arrange into a cubic of 24 monomers. Figure $12$ shows a simple view of a pentagonal dodecahedron, which has 12 equivalent faces. Figure $12$: Pentagonal dodecahedron. 1. Rotating dodecahedron: https://commons.wikimedia.org/wiki/F...decahedron.gif. User Cyp on en.Wikipedia, CC BY-SA 3.0 <http://creativecommons.org/licenses/by-sa/3.0/>, via Wikimedia Commons First let's consider one single E2 monomer. It has a longer disulfide redox domain followed by a smaller dimerization domain which allows the assembly of multiple subunits into the dodecahedron. In greater detail, the monomer has two lipoyl domains, a small domain that allows binding to E1 and a C terminal catalytic domain. Figure $12$ shows an interactive iCn3D model of E2 inner core 60-mer of human pyruvate dehydrogenase (pdb 6CT0). Symmetry axes are not shown Figure $12$: E2 inner core 60-mer of human pyruvate dehydrogenase (pdb 6CT0). (Copyright; author via source). Click the image for a popup or use this external: https://structure.ncbi.nlm.nih.gov/i...zs6pwfKNuJ2VS6 Each of the 60 subunits is shown in light cyan. To see the C symmetry axes: • select the menu = • Choose/Check in order: Analysis, Symmetry, From PDB, 1(global), apply • When you see just a single monomeric chain choose Clear The symmetry axes will then appear. E3: dihydrolipoyl dehydrogenase The sole function of this subunit is reoxidation of the now reduced lipoamide with the free sulfhydryl to the cyclic disulfide form so the enzyme can engage in further catalysis. FAD covalently bound to the E3 subunit is the oxidizing agent. This is our first encounter with FAD. Similarly to NAD+, this dinucleotide gains a hydride (:H-) but also in contrast to NAD+ also a proton to form FADH2. Another way that the FAD/FADH2 differs from NAD+/NADH is that the FAD/FADH2 or their mononucleotide analog (FMN/FMNH2) pairs are either covalently attached (in about 10% of flavoproteins) or bound with such a low KD (often in the nanomolar range) that they don't dissociate from the enzyme during catalysis. Hence after oxidizing a bound substrate, the reduced FADH2 must be reoxidized by another oxidizing agent, often NAD+ which can diffuse into the active site to do its job and then dissociate from the complex in the form of NADH, leaving the enzyme competent for another round of catalysis. (DOI: 10.1002/chem.201704622) Figure $13$ shows an interactive iCn3D model of E3 bound to both FAD (noncovalently) and NADH (NAI) (1ZMD) in the B chain of E3. Symmetry axes are not shown Figure $13$: E3 bound to both FAD (noncovalently) and NADH (NAI) (1ZMD). (Copyright; author via source). Click the image for a popup or use this external: https://structure.ncbi.nlm.nih.gov/i...LAMqAS3Tx9SPE8. Figure $14$ shows an interactive iCn3D model highlighting the noncovalent interactions stabilizing bound FAD and NADH (NAI) in the E3 subunit of pyruvate dehydrogenase (1ZMD). Figure $14$: Noncovalent interactions stabilizing bound FAD and NADH (NAI) in the E3 subunit of pyruvate dehydrogenase (1ZMD). (Copyright; author via source). Click the image for a popup or use this external:https://structure.ncbi.nlm.nih.gov/i...gUED1ZrJVjKcD8 Let's put it all together! Figure $15$ shows a video of the pyruvate dehydrogenase complex from the HHMI. Figure $15$: Video of the pyruvate dehydrogenase complex

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/16%3A_The_Citric_Acid_Cycle/16.01%3A_Production_of_Acetyl-CoA_%28Activated_Acetate%29.txt

Search Fundamentals of Biochemistry Introduction The acetyl-CoA formed by the pyruvate dehydrogenase complex (PDC) now enters a cyclic, non-linear pathway called the citric acid cycle, tricarboxylic acid (TCA) cycle, or the Kreb's cycle after Hans Krebs who discovered it. The cycle is shown in Figure \(1\) in wedge/dash form with stereochemistry included to give a more exact representation. Figure \(1\): Citric Acid Cycle Why is this pathway a cycle and not a linear pathway as we have seen for glycolysis? A simple answer is that it evolved that way, but why would that be advantageous? It turns out that some of the key "intermediates" in the pathway are pulled away for the biosynthesis of other biomolecules. If the citric acid cycle was linear, and intermediates pulled off for other reactions, the linear pathway would taper off, which would not be optimal for a key energy production pathway. Of course, the removal of intermediates from a cyclic pathway would also slow it down but when this happens, enzymes outside of the cycle are used to produce key reaction intermediates of the cycle to it going. The replenishing reactions are called anapleurotic reactions. Krebs, in his detailed analysis of the enzymes involved in "intermediary" metabolism, used radioisotope-labeled reactants to trace carbon atoms in respiring tissue. He found that when radiolabeled pyruvate and oxaloacetate were added to muscle tissue in vitro, radiolabeled citrate was formed. Pyruvate + Oxaloacetate → citrate + CO2 This is correct but omits the initial conversion of pyruvate to acetyl-CoA. Hence the "end product" of the pathway (oxaloacetate) reforms the beginning reactant (citrate) so he surmised that the pathway was circular. The next obvious question is why are eight reactions are required for the oxidation of just 2 Cs in pyruvate. Partly this is a matter of evolution again, as the early evolutionary pathway probably combined an oxidative (clockwise) set of reactions with a reductive (counterclockwise) set. Part of the chemistry in the cycle is devoted to producing either β-keto acids, which are easy to oxidatively decarboxylate, or converting α-ketoacid to molecules with better electron sinks β to the departing CO2. After the net 2 carbon atoms added to the cycle are released as 2 CO2s, the rest of the reactions are used to regenerate oxaloacetate, allowing the cycle to continue. Of course, the ultimate goal of an energy-extractive oxidative pathway is not just to form CO2 but to form ATP or its equivalent (i.e. GTP). Notice that 3 NAD+s are used and converted to 3 NADH. In addition, a new, more potent oxidizing agent, FAD, is used and it is converted to FADH2. NAD+ and FAD are replenished by reoxidation of NADH and FADH2 (reduced forms) back to NAD+ and FAD, through mitochondrial electron transport (oxidation) reactions, in which electrons are passed to stronger and stronger oxidizing agents, the last being O2. In this thermodynamically favored process, lots of ATPs are made. We will explore those reactions in the next section. We will go through each of the steps in the citric acid cycle separately and show how the pathway is regulated (section 16.3). Why such detail? There are only 8 steps. It seems that we should be able to carefully examine each given that the citric acid cycle is a hub that along with glycolysis controls metabolic flow through many interconnected metabolic pathways. At the same time, we can't explore each reaction in every pathway described in this text in great detail, otherwise, this book would become more of an encyclopedia. In this chapter and beyond, we will focus on mechanistic details only of enzymes that catalyze different types of reactions than those in glycolysis or the citric acid cycle, and those with interesting cofactors and mechanisms. Other issues add complexity for learners. The PDC and citric acid cycle reaction occur in the mitochondrial matrix. Cytoplasmic pyruvate and NAD+ must be transported into the matrix from the cytoplasm. In addition, some of the enzymes in the citric acid cycle have both cytoplasmic and mitochondrial variants. Some of these homologous pairs are differentiated by their use of NAD+ or NADP+ as an oxidizing agent. The ones in the cytoplasm are not part of the cycle. You would expect these enzyme pairs to have similar tertiary structures and active site chemistry. Prokaryotic forms of these enzymes are similar structurally to their eukaryotic forms so the interactive molecular models shown below will show enzymes from a variety of organisms. Finally, there are many variants, shunts, and bypasses of the citric acid pathway in different organisms. We will explore this topic in section 16.4 We will try to pair reaction mechanism diagrams that show the flow of electrons in bond-making and breaking with interactive molecular models of the active site. You should rotate the models to align and identify key amino acids and ligands (substrate, substrate analogs, inhibitors, activators) shown in the static 2D mechanism diagrams. Since the active sites are often conserved across prokaryotic and eukaryotic versions, the choice of PDB structures used depends on which best illustrates a conceptual point. Note There are many ways to write abbreviated chemical equations showing NAD+/NADH and FAD/FAD2 and hydrogen ions in metabolic pathway diagrams. To make sense of them, consider a simplified mechanism for the oxidation of ethanol by alcohol dehydrogenase, as shown in Figure \(2\). Note that there are 2 Hs on the oxidized substrate (ethanol) that are involved. One is a hydride and the other is a proton. Figure \(2\): Oxidation of ethanol using NAD+ Here is a list of different and seemingly contradictory ways to write a chemical equation to show changes in NAD+/NADH and H ions: 1. NAD+ + :H- → NADH. This chemical equation is charge balanced and shows just the changes to the NAD+/NADH pair, but it doesn't show the proton (H+) lost from the substrate. 2. NAD+ + 2e- + H+ → NADH. This is the same as equation (1) but with the hydride separated into an electron pair and a proton. 3. NAD+ + H+ → NADH. This is balanced for Hs but not + charge as it doesn't explicitly show the electron pair from the hydride added to NAD+. 4. NAD+ → NADH + H+. This is balanced for charge but not for Hs. The extra H+ is the proton from the oxidized substrate. We will use example 4 above throughout this book. That equation is most useful when trying to account for the change in the number of protons in the individual reactions and entire pathways. We will also write simplified chemical equations involving FAD (in which 2H from the substrate are added) as FAD → FADH2. 1. Citrate Synthase (CS) Oxaloacetate + Acetyl-CoASH + H2O → Citrate + CoASH ΔGo = -7.5 kcal/mol (-31 kJ/mol) Acetyl CoA is a thioester. Hence it is "high energy" compared to its hydrolysis products. (Remember, there is no such thing as a "high energy" bond.) The free energy released in its hydrolysis is used to drive the reaction forward. This is important otherwise citrate would not be formed readily. This reaction feeds the end product of glycolysis into the citric acid cycle. It is summarized in Figure \(3\). Figure \(3\): Summary reaction - Citrate Synthase The enzyme exists in two major conformations, an open and closed form. When the open form, which has a binding site for oxaloacetate, binds the substrate, a shift to the closed conformation forms on the binding site of acetyl-CoA. These changes sequester the bound substrates and exclude water and prevent spurious hydrolysis of acetyl-CoA. The binding occurs sequentially so the kinetics follow a sequential ordered mechanism. Figure \(4\) left shows an animated gif that shows the conformational changes between the citrate-bound version (open, green) and the citrate and CoASH-bound form (blue). The image below right shows a smoother transition between the open and closed form without bound ligands (1cts, 2cts) Figure \(4\): Conformational changes in citrate synthase on binding substrate For a more details view of the enzyme view the Regulation of Citrate Synthase. The mechanism below is from https://chem.libretexts.org/Bookshel...trate_Synthase, with Contributors and Attributions from: In this reaction, a C-C bond must form between the substrates. One way to do that is to make a nucleophilic carbanion ion from the alpha carbon of acetyl CoA. Remember, this is not a decarboxylation reaction so we don't have to worry about an electron "sink" on the beta carbon. Forming the carbanion would be possible since the negative charge on the carbon can be withdrawn to the carbonyl oxygen to form an enolate. The enolate becomes even more stable if the negative oxygen is protonated. So this is reaction is an aldol condensation, the addition of an enolate to an aldehyde or ketone. The carboxylate group of aspartic acid 375 on citrate synthase removes the acidic alpha proton on acetyl CoA, while histidine 274 donates a proton to form the neutral enol, a much more stable molecule than the enolate anion. His 274 continues to stabilize the enol during the reaction. Bound oxaloacetate is stabilized in part by Arg 329. In the next part of the mechanism, a second histidine (320) protonates the carbonyl oxygen of oxaloacetate, activating the carbonyl carbon for nucleophilic attack by the enol in the next step to form (S)-citryl CoA. The hydrolysis of the CoASH occurs when His 320 deprotonates a water molecule, facilitating nucleophile attack on the carbonyl carbon bonded to -SCoA, forming citrate. A plausible mechanism is shown in Figure \(5\). Figure \(5\): Citrate synthase mechanism Figure \(6\) shows an interactive iCn3D model of the pig citrate synthase bound to CoASH and citrate (2CTS) Figure \(6\): pig citrate synthase bound to CoASH and citrate (2CTS). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...WXqcEsu336tcL7 The enzyme is a dimer with monomers shown in different colors. Citrate and CoASH are shown in sticks and labeled. The active site residues are shown as sticks and labeled in each subunit. 2. Aconitase Citrate ↔ Isocitrate ΔGo = +2 (rx 2a), -0.5 (rx 2b) kcal/mol; net ΔGo = + 1.5 kcal/mol (+6.3 kJ/mol) Thinking like a chess player, who must anticipate future moves, the chemical rationale for this reaction is to move an OH to a beta position, which in a subsequent reaction is converted to a beta C=O, so it can act as an electron sink to facilitate decarboxylation in a following reaction! The reaction is readily reversible (note the low ΔGo value) since the reactant and product are simple isomers of each other. Figure \(7\) shows the summary reaction. Figure \(7\): Summary reaction of aconitase The enzyme has an inorganic Fe4S4 cluster. Each Fe in the cluster coordinates to 4 S2- in a cubane structure, but when either citrate or isocitrate is bound, one of the Fe ions interacts with both the Os of a substrate carboxylate shown. The other two carboxylates of isocitrate are stabilized through ion-ion interactions by Arg 446 and 663. Figure \(8\): below shows a plausible partial mechanism. An active site deprotonated serine abstract a proton at the S carbon. This is followed by the formation of the C-C double bond and a release of the resulting cis-aconitate from one bond to the FeS cluster. Figure \(8\): Mechanism of aconitase Figure \(9\) shows an interactive iCn3D model of the bovine S642A aconitase with bound citrate (1C97). Figure \(9\): Bovine S642A aconitase with bound citrate (1C97). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...iJJRGaFoCDnhf7 This cis-aconitate intermediate in the interconversion of citrate and isocitrate must do an 1800 flip around the C=C double bond. This is followed by rehydration to form the other isomer. The deprotonated His 101 abstracts a hydrogen from a water-bound to the FeS cluster, with the hydroxide acting as a nucleophile, which along with the redonation of a hydrogen ion on the protonated Ser 642 the alpha-carbon completes the rehydration step in the formation of the other isomer. Exercise \(1\) Why was the S642A mutant used to produce the structure shown in the above iCn3D model? Answer It allows the binding of substrate/product, in this case, isocitrate, to an inactive enzyme as the active site serine was mutated to a non-nucleophilic alanine of similar size. Hence no bond-making/breaking occurs in the complex. References: https://collab.its.virginia.edu/acce...-_-/index.html 3. Isocitrate Dehydrogenase (IDH) Isocitrate + NAD+ → α-ketoglutarate + NADH + H+ ΔGo = -2.0 kcal/mol (-8.4 kJ/mol) The chemical rationale should be clear. In this step, through an oxidative decarboxylation, the CO2 is removed as NADH is produced. The NADH will be reoxidized back to NAD+ in the electron transport chain, leading to ATP production (see next section). The reaction is summarized in Figure \(10\). Figure \(10\): Summary reaction: isocitrate dehydrogenase Exercise \(1\) Why must the oxidation reaction precede the decarboxylation reaction?. Answer First, a beta-ketoacid intermediate must form, which allows easy decarboxylation of the intermediate as the beta carbonyl provides an electron "sink" to facilitate the decarboxylation. There are two forms of this enzyme (IDH), a cytoplasmic (NADP+) form and a mitochondrial (NAD+ ) form. The cytoplasmic forms from various organisms are homodimers and have a common mechanism of catalysis. In contrast yeast mitochondrial IDH has two subunits, IDH1 (regulatory, binds the allosteric activator citrate and AMP) and IDH2 (catalytic, binds isocitrate and NAD+). Mammalian IDHs are tetramers heterodimers (αβ + αγ), which can also form a heterooctamer (αβ + αγ)2. The alpha chain is the catalytic subunit. Mammalian NAD-IDHs are even more complex than yeast NAD-IDH. These enzymes are composed of three types of subunits, α, β, and γ, which share about 40–52% sequence identity. The α and β form an αβ dimer, while α and γ subunits form αγ. These then interact to form the α2βγ heterotetramer, which effectively forms the holoenzyme. It can also form an active heterooctamer.e. The αγ heterodimer is regulated by citrate and ADP. On citrate binding to the allosteric site, a conformation change occurs to enhance isocitrate binding. ADP enhances the binding of the allosteric regulator citrate. Figure \(11\) shows an interactive iCn3D model of that shows the superposition of the α chains of cytoplasmic IDH (NADP, sky blue, 4L03) and mitochondrial IDH (NAD) (6KDY, salmon). Figure \(11\): Superposition of the A chains of cytoplasmic IDH (4L03) and mitochondrial IDH (NAD) (6KDY) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...zy9yn4NaF9g1c9 Cytoplasmic IDH is sky blue and mitochondrial IDH is salmon. Click the 3 bar menu icon (top left) in the model in the window, scroll down to Alternate, and toggle back and forth between the two forms. The structures of the alpha chains, although not identical, align well. Figure \(12\) shows a probable mechanism for the reaction based on the conserved catalytic site shown in the model above. Figure \(12\): Mechanism of isocitrate dehydrogenase (after https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3706558/) Figure \(13\) shows an interactive iCn3D model of active site of the cytoplasmic human IDH1 in complex with NADP+ and Ca2+/α-ketoglutarate (4L03). Figure \(13\): Active site of the cytoplasmic human IDH1 in complex with NADP+ (NAP1) and Ca2+/ alpha-ketoglutarate (4L03) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...P9DqLoPowEV2d9 Figure \(14\) shows an interactive iCn3D model of the active site of the αβ heterodimer of human IDH3 (6kdy) in complex with NAD+. Figure \(14\): αβ heterodimer of human IDH3 (6kdy) in complex with NAD+(6kdy) . (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...hyyJiMFEqwktB7 The α subunit is shown in gray and the β in cyan. 4. α-ketoglutarate dehydrogenase α-ketoglutarate + NAD+ + CoASH → succinyl CoA + CO2 + NADH + H+ ΔGo = -7.2 kcal/mol (-30 kJ/mol) In the last reaction, α-ketoglutarate was formed. Oh no, you might say! It would have been nice to form a β-ketoacid, which could easily decarboxylation. No worries though. We spent all of section 16.1 explaining the biochemistry used to decarboxylate another α-ketoacid, pyruvate. The same chemistry is used to accomplish the oxidative decarboxlation of α-ketoglutarate. Hence we won't expand on the mechanism here. The reaction is shown in Figure \(15\). Figure \(15\): Summary reaction for α-ketoglutarate dehydrogenase 5. Succinyl-CoA synthetase (SCS) succinyl CoA + GDP + Pi → succinate + CoASH + GTP ΔGo = -0.8 kcal/mol (-3.3 kJ/mol) This is the first step in which the energy change in the cycle is captured specifically in the form of a high energy (with respect to its hydrolysis product) phosphoanhydride bond in the form of GTP (and ATP in some organisms). From a chemical step, the cleavage of the thermodynamically unstable thioester is coupled to the endergonic synthesis of GTP. This can transfer its terminal phosphate to ADP to make ATP in reaction that has a ΔGo of about 0 kcal/mol. The reaction is shown in Figure \(16\). Figure \(16\): Summary reaction for succinyl-CoA synthetase Succinyl-CoA synthases have two subunits, α and β. The enzyme in E. coli is a tetramer (α2β2) with the catalysis occurring at the αβ interface. The alpha-subunits interact only with the beta-subunits, whereas the beta-subunits interact to form the dimer of alpha beta-dimers with CoA bound in each α subunit to a nucleotide-binding loop. Two histidines, His 246 and His 142 are involved in the reaction, with His 246 becoming phosphorylated to form an intermediate in the reaction. A mutation of His 142 to an asparagine (H142N) essentially abolishes enzyme activity. Different SCSs have different specificities for purine nucleoside triphosphates. Organisms, including mammals, may have two different isoforms, one that binds ADP and one that uses GDP (as shown in most diagrams of the citric acid cycle). In E. Coli, the α subunit binds CoASH and contains His 246, which gets phosphorylated. The β subunit determines the specificity for either GTP or ATP. In E.Coli the ATP binding site (Site II "in the ATP-grasp fold") is quite distant from the CoASH site (Site II) so phospho-His 246 must move between the sites in the dimer interface. The three steps in the reaction are shown below, where E is the free enzyme, a . indicates a noncovalent complex, and a - represents a covalent bond (after Biochemistry 2002, 41, 537-546) 1. E + succinyl-CoA + Pi ↔ E . succinyl-PO3 + CoASH 2. E . succinyl-PO3 ↔ E-PO3 + succinate 3. E-PO3 + NDP ↔ E + NTP Figure \(17\) shows an abbreviated mechanism that shows only the involvement of His 246. Figure \(17\): Abbreviated mechanism for succinyl-CoA synthase Kinetic analysis suggests that the three substrates bind in a specific order, catalysis occurs, and then the three products leave. This type of reaction is called an ordered ter ter reaction, and is shown in Figure \(18\). Figure \(18\): Ordered ter ter reaction for succinyl-CoA synthase The iCn3D model below shows key alpha-chain residues in the active site, including the phosphorylated His 246 and bound CoASH. (use iCn3D to visualized 1CQJ, the nonphosphorylated form) I) Figure \(19\) shows an interactive iCn3D model of the complex of ADP and Mg2+ with Dephosphorylated E. Coli Succinyl-CoA Synthetase (1CQI) Figure \(19\): Complex of ADP and Mg2+ with Dephosphorylated E. Coli Succinyl-CoA Synthetase (1CQI). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...BssxsabV1S9cz6 Just one αβ dimer is shown. The α subunit is shown in gray and the β in cyan. Bound Pi and CoASH are labeled. The active site His246 in the α chain is shown in stick and labeled. Figure \(20\) shows an interactive iCn3D model of the active site of pig GTP-specific succinyl-CoA synthetase in complex with succinate and CoASH (5CAE). Figure \(20\): Active site of pig GTP-specific succinyl-CoA synthetase in complex with succinate and CoASH (5CAE). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...HjH9ER5wLqb1T7 6. Succinate Dehydrogenase succinate + FAD ↔ fumarate + FADH2 ΔGo = 0 kcal/mol The enzyme is yet another step in closing the cycle to reform oxaloacetate. It is also our first encounter with FAD as an oxidizing agent. Its reduction product, FADH2, will be reoxidized in the electron transport chain (mitochondrial inner membrane for eukaryotes), producing energy for ATP. Hence it can be considered a proxy ATP-generating reaction. The reaction is shown in Figure \(21\). Figure \(21\): Summary reactoin for succinate dehydrogenase The succinate dehydrogenase enzyme is part of the larger Complex II of the electron transport chain. Complex II has many cofactors involved in its overall activity. It also uses an iron/sulfur cluster cofactor, similar to aconitase, which also produces a C=C doubled bonded intermediate. We will discuss it in greater detail in the chapter on electron transport. For now, let's concentrate on this new cofactor and oxidizing agent, FAD. Many enzymes use FAD/FADH2 in redox chemistry. In contrast to NAD+/NADH, the FAD/FADH2 pair stays tightly bound to the enzyme and doesn't readily dissociate. This means that after one cycle of the enzyme (after FAD is converted to FADH2), the enzyme is functionally "dead". Another oxidizing agent must bind to the enzyme and reoxidize FADH2 back to FAD. The dissociation constants for FAD/FADH2 and its protein binder in a flavoprotein are often in the nanomolar range. In around 10% of flavoproteins, FAD/FADH2 are usally covalently bonded to the enzyme. Figure \(22\) shows an interactive iCn3D model of the Avian respiratory complex II FAD binding subunit with FAD and a malate-like intermediate (1YQ3). Figure \(22\): Avian respiratory complex II FAD binding subunit with FAD and a malate-like intermediate (1YQ3). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...SGac3NuN4XBWk7 Note how buried the FAD is in the middle of the dehydrogenase subunit. The malate-like intermediate (TEO) is shown next to the FAD in yellow spacefill. Exercise \(1\) Succinate dehydrogenase is irreversibly inhibited by the toxin 3-nitroproprionic acid (3np) made by some plants and fungi. Eating moldy sugar cane has led to reported deaths. 1. Draw the Lewis structures of succinate and 3-nitroproprionic acid. Compare them and the total number of valence electrons in each. 2. Here is a link to an iCn3D model showing the interaction of 3np with the enzyme. Explain the mode of action of the toxin. https://structure.ncbi.nlm.nih.gov/i...cja9VKU2ZMCac9 Answer These molecules are structurally similar and isoelectronic (the same number of electrons in their Lewis structures. The inhibitor 3np form a covalent adduct through the guanidino group of Arg 297. This is a key catalytic residue that act as a general base that accepts a proton from succinate in the reaction. Figure \(23\) (top) shows a very general and abbreviated mechanism for the enzyme for the immediate reaction of succinate with FAD. The bottom part of the image shows the amino acids surrounding succinate in the avian (bird) version of the enzyme (pdb 1yq4) Figure \(23\): Top - a very general and abbreviated mechanism for the enzyme for the immediate reaction of succinate with FAD. Bottom - amino acids surrounding succinate in the avian (bird) version of the enzyme (1yq4) Exercise \(1\) One of the amino acids surrounding succinate in the figure above acts as a general base and abstracts a protein from succinate as a hydride is transferred (from a plane above) to FAD. Go to this iCn3D of the active site bound to FAD. Which amino acid is the likely general base? (Note: the figure below shows general protonated states of side chains and not necessarily those involved in the proton abstraction. Go to Analysis, Distance and Distance between 2 atoms to find the likely general base. Answer Arg 297 7. fumarase fumarate + H2O ↔ L-malate ΔGo = -0.9 kcal/mol (-3.8 kJ/mol) The chemical rationale for this reaction is clear - it is the penultimate step in the resynthesis of oxaloacetate, one of the reactants that starts the cycle, allowing the cycle to continue. This reaction introduces an O by hydration which can be oxidized in next step to produce NADH for e- transport/ATP production. The reaction is shown in Figure \(24\). Figure \(24\): Summary reaction for fumarase There are Class I (dimers containing an unstable FeS cluster, examples A and B) and Class II (tetramer, no bound iron, oxygen stable, example C) fumarases. Humans have both cytoplasmic and mitochondrial type II fumarases, resulting from alternative transcription of the fumarase genes. We will consider the type II, fumarase C from E. Coli in the following discussion. The tetramer contains just alpha helices and random coils and has two distinct binding sites. Site A appears to be the active site and contains a buried water molecule. Site A, formed from three of the subunits, binds competitive inhibitors such as citrate and β-(trimethylsilyl)maleate, a cis substrate for fumarase, and is buried. 12 Angstroms away is site B, which is found in only one of the subunits near a pi-helix (H129 through N135) and is more surface-exposed. Each site has a histidine, but mutation of only one H188N in the A site disrupts enzyme activity. Both sites bind multi-carboxylates. The role of site B is a bit unclear, but it is most likely an allosteric site involved in the transfer of product (malate) from the buried site to the surface for ultimate dissociation. Figure \(25\) shows an interactive iCn3D model of the fumarase with beta-(trimethylsilyl)maleate and citrate (1fuq). Figure \(25\): Fumarase with beta-(trimethylsilyl)maleate and citrate (1fuq). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...MewEQFebi4VL48 The monomers in the tetramer are shown in different colors. Citrate (Cit) and beta-(trimethylsilyl)maleate (SIF) are shown in spacefill. Figure \(26\) shows an interactive iCn3D model of the binding site of citrate, a competitive inhibitor, of fumarase (1FUQ). Figure \(26\): Binding site of the competitive inhibitor citrate in fumarase (1FUQ). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...yfPc5iPWbExr86 Figure \(27\) shows a plausible mechanism for the trans addition of water to fumarate. Figure \(27\): Mechanism for fumarase 8. Malate Dehydrogenase (MDH) L-malate+ NAD+ ↔ oxaloacetate + NADH + H+ ΔGo = +7.1 kcal/mol (30 kJ/mol) We are finally there! This last reaction of the citric acid cycle produces oxaloacetate, the starting reactant, so the cycle can continue. It also produces NADH for mitochondrial e- transport/ATP production. Notice that is thermodynamically unfavorable (in the standard state) but the reaction is pulled to citrate formation by the first and next step of the cycle, citrate synthase. The reaction is shown in Figure \(28\). Figure \(28\): Summary reaction for malate dehydrogenase Malate dehydrogenases are found in the cytoplasm, where it is part of the aspartate-malate shuttle that moves cytoplasmic malate (and through MDH indirectly NADH) into the mitochondria.  It is also found in the mitochondria, where it is part of the citric acid cycle. There are also NAD+ and NADP+-dependent forms. Malate can undergo two different types of oxidation reactions, one producing oxaloacetate and using NAD+, and one, an oxidative decarboxylation producing pyruvate and CO2, using NADP+. The latter is sometimes called malic enzyme. Humans have two forms (MDH 1 and MDH 2) that use NAD+. The enzyme is a homodimer in humans with binding sites on both. It activity is allosterically regulated by citrate, and it is inhibited by many things, including ATP, ADP, AMP, fumarate, citrate, aspartate, and high concentrations of oxaloacetate. The enzyme is similar to lactate dehydrogenase, which we encountered in the chapter of glycolysis. Kinetic analyses show that NAD+ binds first followed by malate. Figure \(29\) shows an interactive iCn3D model of NAD+ and malate bound to human malate dehydrogenase 2 (4wlu). Figure \(29\): NAD+ and malate bound to human malate dehydrogenase 2 (4wlu). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...DrAXBdu64GPzx8 Just one monomer is shown. NAD+ and malate (LMR) are shown in sticks and labeled. Figure \(30\) shows an abbreviated mechanism for malate dehydrogenase. The numbers refers to the E. Coli enzyme. Figure \(30\): Abbreviated mechanism for malate dehydrogenase Note that the hydride transferred from the malate is shown in red as a deuterium (D). It is transferred to the re face of NAD+ to form NADH. The carbon with the transferred deuterium in NADH is prochiral. Think of that as that carbon being chiral if one of the 2 Hs could be arbitrarily assigned a higher priority in assigning R/S isomers. D has a higher priority than H in the Cahn/Ingold designation system. In the reverse reaction, the D atom, which is above the plane of the ring, occupies the proR position. The proR deuterium is transferred back in this reversible reaction. A D was used simply to indicate the stereochemistry and to assign it in NADH to the proR position. Figure \(31\) shows an interactive iCn3D model of the active site of the E. Coli malate dehydrogenase with bound citrate and NAD+ (1EMD). Figure \(31\): Active site of the E. Coli malate dehydrogenase with bound citrate and NAD+ (1EMD). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...mHhmeFQyPpZj18 We have shown many renderings of the enzymes involved in the cycle. Yet another one is shown below. Figure \(32\) shows an interactive iCn3D model of the electrostatic surface potential of malate dehydrogenase (4WLU). Figure \(32\): Electrostatic surface potential of malate dehydrogenase (4WLU)with bound NAD+ (4WLU). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?izf2dsWXTw2bE8YD7 The display surface is the electrostatic surface potential map of the enzyme. Red shows the surfaces that are more anionic and with a negative electrostatic potential to which cationic molecules would be attracted, while blue represents more cationic surfaces to which the anion would be attracted. Note that the bound NAD+, which has many oxygens which are slightly or fully negative, is bound in a blue, positive electrostatic potential region. Summary Let's do some stoichiometry for the full cycle. Here is the net reaction (assuming that the GTP produced by succinyl-CoA synthetase is equivalent to 1 ATP). Acetyl-CoA + 3NAD+ + FAD + ADP + Pi + 2H2O → 2CO2 + 3NADH + FADH2 + ATP + 2H+ + CoASH This must seem like a lot of work to produce just 1 ATP, especially since the partial, anaerobic oxidation of glucose in glycolysis produced in net fashion 2 ATPs. The key, however, is to realize that 3 NADHs and 1 FADH2 are produced, which when they are reoxidized in mitochondrial electron transport/oxidative phosphorylation, will produce multitudes of ATP. As was true for glycolysis, this main energy-extracting pathway is highly regulated. We will this in the next section. Figure \(33\) shows some key points about each reaction in the citric acid cycle. Figure \(33\): Summary of the citric acid cycle.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/16%3A_The_Citric_Acid_Cycle/16.02%3A__Reactions_of_the_Citric_Acid_Cycle.txt

Search Fundamentals of Biochemistry Overview Entry of pyruvate into the citric acid cycle leading to the aerobic production of energy and intermediates for biosynthesis is a key metabolic step. Hence both the pyruvate dehydrogenase complex and key enzymes in the cycle are targets for regulation. This occurs through substrate availability, product inhibition, allosteric effectors, and post-translational modifications of key enzymes in the pathway. A summary figure showing key regulators is shown in Figure \(1\). Figure \(1\): Regulation of the citric acid cycle https://www.nature.com/articles/s41598-021-98314-z Note that the key steps are regulated mainly by the ratio of mitochondrial NAD+/NADH which is highly influenced by the ratio of ATP/ADP. High NADH inhibits the regulatory enzymes. High levels of acetyl CoA, derived from pyruvate dehydrogenase and also through fatty acids catabolism increases flux through the cycle in part by allosterically activating the first enzyme in the pathway, citrate synthase. The material below is derived from Renée LeClair, Ph.D., Cell Biology, Genetics, and Biochemistry for Pre-Clinical Students. https://med.libretexts.org/@go/page/37584. openly licensed (CC BY-NC-SA 4.0 Regulation of the pyruvate dehydrogenase complex (PDC) Under aerobic conditions, the pyruvate produced by glycolysis will be oxidized to acetyl CoA using the pyruvate dehydrogenase complex (PDC) in the mitochondria (note: its genes are encoded in the nucleus). As this enzyme is a key transition point (the gatekeeper) between cytosolic and mitochondrial metabolism and is highly exergonic (ΔG0' = -7.9 kcal/mol, -38 kJ/mol), it is highly regulated by both covalent and allosteric regulation. Deficiencies of the PDC are X-linked and present with symptoms of lactic acidosis after consuming a meal high in carbohydrates. This metabolic deficiency can be overcome by delivering a ketogenic diet and bypassing glycolysis altogether. The PDC is regulated by allosteric and covalent regulations. The complex itself can be allosterically activated by pyruvate and NAD+. Elevation of the substrate (pyruvate) will enhance flux through this enzyme as will the indication of low energy states as triggered by high NAD+ levels. The PDC is also inhibited by acetyl CoA and NADH directly. Product inhibition is a very common regulatory mechanism and high NADH would signal sufficient energy levels, therefore decreasing the activity of the PDC. Figure \(2\) summarizes the regulation. (Adapted from Marks’ Medical Biochemistry) Figure \(2\): Regulation of pyruvate dehydrogenase. The PDC is also regulated through covalent modification. Phosphorylation of the E1 subunits of the complex will decrease the activity of the enzyme. The enzyme responsible for the phosphorylation of the PDC is pyruvate dehydrogenase kinase. The kinase is regulated inversely to the PDC, as shown in Figure 1 above. The kinase is most active when acetyl-CoA, NADH, and ATP are high. These compounds will stimulate the kinase to phosphorylate and inactivate the PDC. PDK is inhibited by dichloroacetate, TPP, Ca2+, and pyruvate. The PDC can be dephosphorylated by a calcium-mediated phosphatase, PDP. Starvation and diabetes result in increased phosphorylation and inhibition of the complex, which impairs glucose oxidation. Phosphorylation occurs on Serine 264 of the α subunit (site 1), Ser271 (site 2), and Ser203 (site 3) are located on a conserved phosphorylation loop. Sites 1 and 2 (in loop A) are involved in the stabilization of TPP in the active site, while Ser 203 in the adjacent loop B binds Mg2+ which stabilizes PP on bound TPP. All it takes for inhibition is the phosphorylation of just one of the Ser side chains. Phosphorylation prevents the ordering of the loop which occurs on TPP binding which hinders the binding of the lipoyl domains of the PDC core to E1p, which inhibits the flow of metabolites in the PDC. Prevention of PDC phosphorylation by the specific PDK inhibitor dichloroacetate increases levels of reactive oxygen species in mitochondria, which promotes the expression of a mitochondria-K+ channel axis, leading to cellular apoptosis and the inhibition of tumor growth A summary of pathway regulation is shown in Figure \(3\). Figure \(3\): Summary of the regulation of pyruvate dehydrogenase In general, PDC is activated through its substrates CoASH and NAD+, and kinase inhibition or phosphatase activation (PDP) (dichloroacetate, TPP, Ca2+, and pyruvate), is inhibited by its products acetyl CoA and NADH and activation of kinase (PDK). Abbreviations : PDC: pyruvate dehydrogenase complex, PDK: pyruvate dehydrogenase kinase, PDP: pyruvate dehydrogenase phosphatase TPP: thiamine pyrophosphate. The complex is also acetylated and succinylated. Table \(1\) below shows a summary of the regulation of pyruvate through glycolytic enzymes and pyruvate dehydrogenase Metabolic Pathway Major Regulatory Enzyme(s) Allosteric Effectors Post-translational modifications Hormonal Effects Glycolysis hexokinase; glucokinase (liver) Glucose 6P (-) PFK-1 Fructose 2,6BP, AMP (+) Citrate (-) ↑ Insulin/Glucagon leads to dephosphorylation of PFK2 and increases producthe tion of F2,6BP Pyruvate Kinase Fructose 1,6BP (+) ATP, Alanine (-) ↑ Insulin/Glucagon leads to dephosporylation Pyruvate Dehydrogenase PDC Pyruvate, NAD+ (+) Acetyl CoA, NADH, ATP (-) dephosphorylation by PDP (+) phosphorylation by PDK (-) ↑ Insulin/Glucagon leads to dephosphorylation Table \(1\): Summary of the regulation of pyruvate through glycolytic enzymes and pyruvate dehydrogenase Regulation of Citrate Synthase Citrate synthase (ΔGo = -7.5 kcal/mol, -31 kJ/mo), Isocitrate dehydrogenase (ΔGo = -2.0 kcal/mol, -8.4 kJ/mol) and alpha-ketoglutarate dehydrogenase (ΔGo = -7.2 kcal/mol, -30 kJ/mol) are all exergonic and likely candidates for regulation. Indeed they are. Let's start with citrate synthase. Citrate synthase is regulated in part by the availability of substrate acetyl-CoA and oxaloacetate. It is inhibited by NADH and also citrate, a competitive inhibitor of oxaloacetate binding. Succinyl-CoA, a downstream product of the citric acid cycle, is a competitive inhibitor of acetyl-CoA binding. Figure \(4\) shows an interactive iCn3D model of a structural comparison of pig citrate synthase with bound citrate (1CTS) and with bound citrate and CoASH (2CTS) Figure \(4\): Structural comparison of pig citrate synthase with bound citrate (1CTS) and with bound citrate and CoASH (2CTS). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...94UeRDsFyGTcdA Toggle the "a" key back and forth to change from the open structure (gray) with bound citrate (1CTS) to the closed structure (cyan) after CoASH binds (2CTS). Citrate and CoASH are the products of citrate synthase reaction, but seeing how they interact with the protein gives clues into catalysis. When both are bound, the enzyme is in closed conformation which would prevent spurious hydrolysis of the actual acetyl-CoA when the reaction proceeds to citrate formation. In just the presence of citrate, the enzyme is in open form, allowing the release of citrate as a product. The binding of the reactant oxaloacetate triggers the conversion to the closed form. NADH is reported to be an allosteric inhibitor of bacterial citrate synthases but no entries are available for the binding of NADH to mammalian enzymes. The effects of binding acetyl-CoA on the structure of citrate synthase are shown in Figure \(5\): Figure \(5\): Effect of acetyl-CoA binding on CS structure. Omini, J., Wojciechowska, I., Skirycz, A. et al. The association of the malate dehydrogenase-citrate synthase metabolon is modulated by intermediates of the Krebs tricarboxylic acid cycle. Sci Rep 11, 18770 (2021). https://doi.org/10.1038/s41598-021-98314-z. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/. Panel (A) shows the open format of CS (PDB ID, 1cts) in a cartoon model, with cylindrical α-helices containing a citrate molecule in a stick model. Subunits A and B are colored white and cyan, respectively. Key residues, A266Lys, A46Arg, and B164Arg, are shown in the sticks’ side chains. The residues are shown in the order of the chain name, residue number, and amino acid. the dimer structure was generated using crystallographic symmetry. One active site domain composed mainly of the A-chain is shown. Panel (B) shows the superimposed model between the open (1cts; white) and close (2cts; wheat) formats. The citrate molecule is at the same location with a slight rotation. The CoA molecule is present only in the closed format. The molecular domain shown can be divided into the movable upper half and the rigid bottom half. In the bottom domain, A45Arg and B146Arg are shown in the stick model. The locations of those two Cα in Arg are almost consistent between the open and closed formats. The top half domain is movable. The motion is visible as the rotation of α-helices represented by A312Gly and A365Gly, indicated by arrows. The A366Lys moves inward and forms a hydrogen bond network A366Lys (NZ):: A438COA(O8A):: B164Arg(NH1). Panel (C) shows the closed format of CS (PDB ID, 2cts) with citrate and CoA molecules in stick models. Panel (D) shows the surface electrostatic potential of the open format CS excluding ligands. Calculations were performed in the vacuum environment and ranged between − 71 and + 71. Red and blue represent the negative and positive potentials. The domain shown corresponds to panel A. Patches of negative charge (NC) and hydrophobic area (HF) are observed. Panel (E) shows the surface electrostatic potential of the closed format CS excluding ligands. The domain shown corresponds to panel D. Patches of positive charge (PC1, PC2) are observed. We will see how the electrostatic surface potential of citrate synthase allows it to interact with other citric cycle enzymes to form a metabolon at the end of this chapter section. Regulation of Isocitrate Dehydrogenase In the previous section, structures of the αγ and αβ heterodimer building blocks of the protein were described. The α subunits contain the catalytic site while the β and γ subunits were the regulatory subunits that bind allosteric effectors. Citrate and ADP allosterically activate both the α2βγ heterotetramer and αγ heterodimer. They bind next to each other in the allosteric site, along with Mg2+. Conformational changes in binding citrate lead to a change in and activation of the catalytic subunit α. The binding of ADP just enhances this effect. The domain and cartoon structure of the αγ heterodimer of human NAD-IDH are shown in Figure \(6\). Figure \(6\): Domain and cartoon structure of the αγ heterodimer of human NAD-IDH. Ma, T., Peng, Y., Huang, W. et al. Molecular mechanism of the allosteric regulation of the αγ heterodimer of human NAD-dependent isocitrate dehydrogenase. Sci Rep 7, 40921 (2017). https://doi.org/10.1038/srep40921. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ The top panel shows the domain structure of the two monomers. The bottom panel shows two views of the dimer. The color coding is the same as in the top panel. Figure \(7\) shows bound citrate and ADP in the allosteric binding site in the γ subunit of IDH. Figure \(7\): Bound citrate and ADP in the allosteric binding site in the γ subunit of IDH. Ma et al, ibid. The color represents the electrostatic surface potential of the site with blue indicating more positive and red more negative. Note that both allosteric activators bind adjacent to each other. The binding of ADP does not change the conformation after the citrate is bound. The proposed molecular mechanism for allosteric regulation of IDH is shown in Figure \(8\). Figure \(8\): Mechanism mechanism of allosteric regulation of the αγ heterodimer of IDH. Ma et al, ibid. Legend: In the absence of any activators, the active site adopts an inactive conformation unfavorable for the ICT binding, and the enzyme is in the basal state which has a high S0.5,ICT with a low catalytic efficiency. The binding of CIT induces conformational changes at the allosteric site, which are transmitted to the active site through conformational changes of the structural elements at the heterodimer interface, including the β5–β6 loop, the α7 helix, and the β7-strand in both the α and γ subunits, leading to the conversion of the active site from the inactive conformation to the active conformation favorable for the ICT binding. Hence, the enzyme assumes the partially activated state which has a moderately decreased S0.5,ICT (lower substrate concentration for half-maximal activity) with a moderately increased catalytic efficiency. The binding of ADP in the presence of CIT does not induce further conformational changes at the allosteric site and the active site but establishes a more extensive hydrogen-bonding network among CIT, ADP, and the surrounding residues through the metal ion, which conversely enhances or stabilizes the CIT binding. Hence, the binding of CIT and ADP together has a synergistic activation effect, and the enzyme assumes the fully activated state which has a substantially decreased S0.5,ICT with a significantly increased catalytic efficiency. Regulation of α-ketoglutarate dehydrogenase α-ketoglutarate dehydrogenase and pyruvate dehydrogenase complex both catalyzed the oxidative decarboxylation of α-ketoacids. They use a common mechanism involving three enzymes, E1-E3, in a large complex. The regulation of α-ketoglutarate dehydrogenase activity is shown in Figure \(9\). Figure \(9\): Regulation mechanisms of α-ketoglutarate dehydrogenase complex (α-KGDC). Vatrinet, R., Leone, G., De Luise, M. et al. The α-ketoglutarate dehydrogenase complex in cancer metabolic plasticity. LS and DHLA are lipoamide and dihydrolipoamide, respectively. TPP is thiamine pyrophosphate. Cancer Metab 5, 3 (2017). https://doi.org/10.1186/s40170-017-0165-0. Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), As with the other key regulatory enzymes, α-KGDC is regulated by ATP/ADP and NADH/NAD+ ratios. The product, succinyl-CoA inhibits the reaction at E2. The mitochondria are reservoirs for Ca2+ ions. These ions increase the activity of pyruvate, isocitrate, and α-ketoglutarate dehydrogenases with α-ketoglutarate dehydrogenases most affected. Calcium effects also depend on ATP/ADP and NADH/NAD+ ratios. Regulation by Metabolon Formation Several citric acid cycle enzymes interact to form a metabolon, which allows enhance flux through pathways as substrate and products are channeled directly from one enzyme to another in the complex. This "facilitated" diffusion minimizes the dissociation of substrates/products and enhances catalysis. Three enzymes in the citric acid cycle, citrate synthase, malate dehydrogenase, and aconitase form a metabolon, as shown by chemical cross-linking and docking studies. The linkages between malate dehydrogenase, which produces oxaloacetate, and citrate synthase, which uses it, are important since concentrations of oxaloacetate are low and would produce reduced flux in the citric acid cycle if it were not channeled directly into citrate synthase. Malate dehydrogenase is also not favored to produce oxaloacetate based on standard free energy and Keq values so pulling the reaction towards citrate synthase in the metabolon also helps the flux. L-malate+ NAD+ ↔ oxaloacetate + NADH + H+ ΔGo = +7.1 kcal/mol (+30 kJ/mol) Figure \(10\) shows models of malate dehydrogenase (MDH) and the open and closed forms of citrate synthase (CS). Electrostatic interactions are key. Figure \(10\): Models of malate dehydrogenase (MDH) and the open and closed forms of citrate synthase (CS). Omini, J., Wojciechowska, I., Skirycz, A. et al. Association of the malate dehydrogenase-citrate synthase metabolon is modulated by intermediates of the Krebs tricarboxylic acid cycle. Sci Rep 11, 18770 (2021). https://doi.org/10.1038/s41598-021-98314-z. http://creativecommons.org/licenses/by/4.0/.Creative Commons Attribution 4.0 International License. Panels (F) and (G) show simulated interactions between MDH (green) and CS in its open (white) form (panel F) and between MDH (green) and CS in its closed (wheat) format (panel G). The 65Arg and 67Arg residues that are involved in the MDH-CS interaction are highlighted in blue. Active site residues, His 274, His320 (blue), and Asp375 (red) were shown. Panel (H) Predicted acetyl-CoA binding sites in CS apoenzyme. The white surface model of the CS apoenzyme in the open format is shown. The 274His, 320His, 65Arg, and 67Arg residues are highlighted in blue. The orange mesh indicates the positions of acetyl-CoA at the predicted binding sites. White and orange stick models indicate the citrate and CoA in the reported crystal structure, respectively. Does the activity of citrate synthase change when it is complexed to malate dehydrogenase in a metabolon? The results of studies probing that question show that it is affected. The results of such studies are shows that it does as illustrated in Figure \(11\). Figure \(11\): Effects of metabolites involved in the MDH and CS reactions on the affinity of the MDH-CS multi-enzyme complex. Omini et al, ibid. Curves represent the response (fraction bound) against CS concentration (M). The interaction was assessed in the MST buffer (control, green) or those with 10 mM of metabolites. Error bars represent the standard deviations of three measurements. Asterisks indicate the conditions that showed significant Kd differences with no Kd confidence overlap with the control. Panel (A) shows the effects of the reaction substrates of MDH and CS. The MDH-CS interaction was assessed in the presence of acetyl-CoA (red), NAD+ (blue), or malate (brown). Panel (B) shows the effects of the reaction on products of CS and MDH. The MDH-CS interaction was assessed in the presence of CoA (grey), NADH (orange), or citrate (green). Panel (C) shows the effects of oxaloacetate (OAA) in combination with other CS substrates. The effects of sole substrates, acetyl-CoA (red), OAA (blue), and NADH (orange), as well as their combinations, OAA/acetyl-CoA (purple) and OAA/NADH (gray), were analyzed. These curves clearly show how the ratio of NADH/NAD+ affects the activity of citrate synthase when it is part of a metabolon. NAD+ increases activity (Panel A) while NADH decreases it (Panel B).

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/16%3A_The_Citric_Acid_Cycle/16.03%3A_Regulation_of_the_Citric_Acid_Cycle.txt

Search Fundamentals of Biochemistry Citric Acid Cycle Shunts and Bypasses Evolution has allowed variants in the citric acid cycle to produce new functionalities in organisms. Let's consider a few. Glyoxylate Shunt What if you were a microorganism that has evolved to use acetate (2C) as a source (if not the sole source) of energy? Remember that the citric acid cycle is also used to generate metabolites for anaplerotic reactions (from the Greek ἀνά= 'up' and πληρόω= 'to fill'). In the full citric acid cycle, activated acetate (2C) reacts with oxaloacetate (4) to produce the 6C molecule citric acid. As the cycle continues back to oxaloacetate, 2 Cs are lost as CO2, so in sum for 1 turn of the cycle: oxaloacetate (4C) + acetyl-CoA (2C) → Citrate (6C) → → → succinate (4C) + 2 CO2 (2C). In the process, oxidation reactions occur leading to the formation of NADH and FADH2 and the metabolites formed can be withdrawn for biosynthesis (OAA, αKG, and succinyl CoA). An alternative "cycle" would be to eliminate the two decarboxylation reactions and generate a 4C molecule (succinate) and one 2C metabolite (glyoxylate), which could react with another activated acetate (acetyl-CoA) to form oxaloacetate (or more precisely malate which can be oxidized to it). Hence two acetates are required, but if you are organisms adapted to use this as a metabolic energy source, it's no problem. Here is the next reaction oxaloacetate (4C) + acetyl-CoA (2C) → Citrate (6C) → → succinate (4C) + glyoxylate (2C); glyoxylate (2C) + acetyl-CoA (2C) → malate (4C ) This reaction would act as a metabolic shunt, altering the flow of metabolites by bypassing part of the citric acid cycle. The shunt, which is found in many microorganisms and some plants is called the glyoxylate shunt, which is shown in Figure \(1\). Why is this so cool? First, consider this observation about the citric acid cycle. One acetate, in the form of acetyl-CoA, is needed to form one oxaloacetate in one turn of the cycle, which in turn is needed in the next turn of the cycle. Hence acetyl-CoA cannot, in a net fashion, be used to synthesize oxaloacetate. We saw previously that oxalate is an intermediate in the synthesis of phosphoenolpyruvate from pyruvate in gluconeogenesis. Hence acetate can not be used to form glucose in a net fashion. (Note: pyruvate dehydrogenase is not reversible). Another way to think about this is 2Cs enter the cycle as acetyl-CoA and 2 leave as CO2 so no net synthesis can occur. And, you are in trouble if you draw off oxaloacetate as well as α-ketoglutarate and succinate for anaplerotic reactions for biosynthesis as they are needed metabolites for a cycle that consumes and produces one of each of these will be depleted if withdrawn Now if you are an organism (many bacteria and plants) that has the glyoxylate shunt, you have no worries. A mini cycle still occurs with the conversion of isocitrate to glyoxylate to malate and back around to oxaloacetate and citrate (for energy production) with the net production of 1 succinate (for biosynthesis). The only Cs "lost" from the mini cycle are in the form of succinate so they are not lost as they are as 2 CO2s in the citric acid cycle since they are used for biosynthesis. The next reaction for the glyoxylate cycle is: 2 Acetyl-CoA + 2 NAD+ + 2 H2O → 2 succinate + 2 NADH + 2 CoASH + 2 H+ . In plants, this reaction occurs in organelles called Glyoxysomes. The succinate can be converted through its continuation through the citric acid to oxaloacetate which can then form glucose and other carbohydrates through gluconeogenesis. Hence organisms that have the glyoxylate shunt (or cycle) can synthesize carbohydrates in a net fashion from acetate, which can derive from fatty acid degradation as we will see in a subsequent chapter. Now to operate the glyoxylate cycle efficiently, you do need to activate the carboxyl end of acetate. Plants and other organisms that can grow on acetate can produce acetyl-CoA needed for the above pathways from the following reaction catalyzed by acetyl-CoA synthase (i.e. not pyruvate dehydrogenase) Acetate + CoASH + ATP → Acetyl-CoA + AMP + PPi Note that one phosphoanhydride bond is cleaved and one thioester bond is formed. These molecules are both high energy compared to their hydrolysis products so this reaction alone is not thermodynamically favored in any significant way (remember that there is no such thing as a high energy bond).  The hydrolysis of pyrophosphate (PPi) drives this reaction forward. An analogous reaction drives the creation of peptide bonds in protein as that reaction also uses ATP to create an activated mixed anhydride from the free carboxyl end GABA Shunt Now let's move to the brain where there is lots of chemistry happening as you read this page. Relatively high concentrations of key neurotransmitters are required for neural function and these need to be maintained against metabolic losses. Glutamate is the principal excitatory neurotransmitter and GABA (δ-aminobutryic acid) is the principal inhibitory neurotransmitter. Both are maintained at high concentrations and are principal players in the GABA shunt. Figure \(2\) shows a part of the large pathway shown above but in a more expanded form. GABA is formed in the cytoplasm by decarboxylation of Glu by the enzyme glutamate decarboxylase, which appears to be expressed predominantly in neural tissue. It is metabolized by GABA transaminase but only if the compound from which the shunt starts, α-ketoglutarate, is present abundantly. This conserves the supply of GABA in the neuron. Stated in another way, the breakdown product of GABA, succinic semialdehyde, is formed only if GABA's precursor is present. Inhibitors of GABA aminotransferase are used to treat epilepsy. Another function of the shunt is that it effectively allows glutamate to loop into the cycle. α-ketoglutarate ↔ Succinate Bypass In anticipation of the section below, one could ask the following question: What happens when a step in the cycle is impaired or missing? Cyanobacteria, a key player in atmospheric O2 production and drawn down of natural and anthropogenic atmospheric CO2, were thought to have an incomplete citric cycle as they lack α-ketoglutarate dehydrogenase. They work around this issue by converting α-ketoglutarate to succinate directly using two enzymes, alpha-ketoglutarate decarboxylase, and succinic semialdehyde dehydrogenase, as shown below. The first enzyme catalyzes a non-oxidative decarboxylation of the substrate to succinic semialdehyde. This bypass is shown in Figure \(3\). In addition, cyanobacteria also use the GABA shunt as a bypass for α-ketoglutarate dehydrogenase. Here is yet another example. Each member of the cycle is an important member. Let's consider the first one, citrate. It is formed in the key step coupling the output of anaerobic metabolism of glucose, pyruvate) with the formation of citrate in the citric acid cycle. What happens if aerobic metabolism is impaired, as it would in hypoxic and anoxic conditions, or if mitochondria function is compromised or in disease states such as cancer? Might there be another way to form citrate? Turns out that there is. and it centers on α-ketoglutarate again. It is illustrated in the above figure. Lets's consider cancer which is characterized by a state of rapid proliferation of cells. This requires both energy and metabolic precursors for the biosynthesis of carbohydrates, lipids, and proteins. The citric acid cycle offers both. To accommodate this demand for the efflux of citric acid cycle intermediates for reductive biosynthesis, large fluxes of glutamate (derived from abundant glutamine) into the cycle are used in the form of α-ketoglutarate. In another anaplerotic reaction, citrate can be cleaved by citrate lyase to form acetyl-CoA which can be used for fatty acid synthesis needed by rapidly proliferating cells. To replenish the citrate, cancer cells convert α-ketoglutarate by reductive carboxylation to isocitrate by isocitrate dehydrogenase 2, which uses NADPH for the reduction reaction. That NADPH comes from the enzyme nicotinamide nucleotide transhydrogenase, which can interconvert matrix pools of NADH and NADPH using the collapse of a protein gradient across the mitochondria inner membrane (which we will study in a subsequent chapter). H+(in) + NAD+ + NADPH ↔ H+(out) + NADH + NADP+ Note that both oxidative (α-ketoglutarate to succinyl CoA) and reductive reactions (α-ketoglutarate to isocitrate) occur in this process. Variants of the TCA cycle in Microorganisms It's very easy to be anthropocentric in constructing a biochemistry text as many who take the course are interested in human medicine. Since a human is an ecosystem of organisms with an expansive microbiome on their skin and in their gut, even from a human health perspective, it is important to compare the same pathway in different organisms. It's also important to understand our role as a small part of a vast biosphere where our survival depends on other organisms, large and small. In that light, let's consider the citric acid cycle of other organisms. It seems that most organisms have the anaerobic and universal glycolytic pathway. How about the aerobic citric acid cycle? These days of single-cell genomic analysis make it simple in principle to analyze the citric acid cycle genes of any organism. Variants of it are found in generally all aerobic organisms and even some anaerobic one. Some subtle differences exist between eukaryotic and prokaryotic organisms. NAD+ is used as a substrate in the mammalian form of isocitrate dehydrogenase while prokaryotes use NADP+. An NAD+-dependent malate dehydrogenase is used in mammals while some prokaryotes use a different enzyme, a NADP+-dependent malate-quinone oxidoreductase. Lastly different enzymes (and unfortunately with varying names) are used to convert succinyl-CoA to succinate. Plants and fungi use ADP as a substrate, mammals have two different enzymes often named different and for the reverse reaction as succinate-CoA ligase (ADP forming) and succinate-CoA ligase (GDP forming). Analyzes shown that few bacteria have complete cycles. Of those with incomplete cycles, the early steps are least conserved, while the latter are most conserved. The cycle is used not only for the oxidative production of energy but also the generation of metabolites (α-ketoglutarate, oxaloacetate, and succinyl-CoA), which are pulled from the cycle for biosynthesis. Autotrophs that don't have a complete cycle can make those products from pyruvate. Consider α-ketoglutarate. Some make it through the clockwise oxidative reaction from citrate to α-ketoglutarate while some methanogenic Archaea make it through counterclockwise reduction reactions. Comparative genomic analyses suggest that the citric acid cycle probably arose from a "linear" oxidative pathways leading to α-ketoglutarate and a reductive one leading to succinyl CoA. Knowing the pathways in individual microorganisms can assist in the rational drug design of new antibiotics. Figure \(4\)below, adapted from Huynen et al, shows a geometric analysis diagram of the citric acid cycle and variants in other organisms. The α-ketoacid pathway - A primordial, prebiotic anabolic "TCA-like" pathway You might be wondering after studying the complexity of pyruvate dehydrogenase and the citric acid cycle how the cycle might have originated. As discussed above, clues come from comparative genome analysis of genes encoding the enzymes for the reactions in a multitude of organisms. By looking at evolutionary changes in genes and functions for these reactions, one can obtain some ideas of the first enzymes that evolved in the pathway. But what about an abiotic pathway that might have arisen before the first biological one? Stubbs et al have shown that the very simple molecules glyoxylate and pyruvate can react in mild aqueous conditions (pH 7 in 0.5 M phosphate buffer heated to 50 °C) to form, in a single pot, α-ketoacids similar to present citric acid cycle intermediates. From a kinetic perspective, they were formed in the reverse, counterclockwise reductive direction compared to the citric acid cycle (clockwise, oxidative). No metals or "enzymes" were needed for the transformations. Glyoxylate acts as a reducing agent and is simultaneously the source of carbon atoms. Once these α-ketoacids form, they could be theoretically converted by transamination reactions to amino acids as they should be characterized by a Keq ≈1 and a ΔG0 0. The citric acid cycle is filled are carboxylic acids. Only the α-ketoacids (α-ketoglutarate and oxaloacetate) can readily form carbanions at the alpha carbons as their Hs are slightly acidic due to resonance stabilization of the carbanion through and enolate tautomer. The others are electron-rich so they are less acidic at the alpha carbons, discouraging deprotonation in the absence of harsh catalysts. So perhaps alternative functional groups were present in abiotic precursors. Perhaps they contained α-ketoacids similar to α-ketoglutarate, oxaloacetate, and of course pyruvate. In addition, α-ketoacids have an electrophile in the form of the carbonyl carbon form C-C bond formation. Figure \(5\) shows the reductive, counter-clockwise pathway they determined. Note that in both the glyoxylate "cycle" which has also been proposed as an abiotic pathway and the α-ketoacid reductive pathway, no CO2 is lost in any step. That suggests that the actual decarboxylation steps, although often favored thermodynamically, required the development of catalysts. The two α-ketoacids substrates in this pathway, glyoxylate (2Cs) and pyruvate (3Cs) are the smallest and most likely formed acids available. Figure \(6\) is an animation that shows the entire citric acid cycle (catabolic, oxidative, running clockwise), the glyoxylate shunt which bypasses many steps in the citric acid cycle, and the proposed prebiotic anabolic alpha-ketoacid pathway (bold green arrows) which essentially runs counterclockwise and is reductive.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/16%3A_The_Citric_Acid_Cycle/16.04%3A_Variants_of_the_Citric_Acid_Cycle.txt

Fatty acid catabolism is the mechanism by which the body accesses energy stored as triglycerides. • 17.1: Digestion, Mobilization, and Transport of Fats In this chapter we will discuss the breakdown of fats to produce useful energy for biosynthesis and for ATP production. • 17.2: Oxidation of Fatty Acids Fatty acids, esterified to glycerol in triacylglycerols, are the major source of stored energy in organisms. As we burn fossil fuels to produce energy to drive our society, so can we "burn" fatty acids to produce energy for heat, to drive biosynthetic reactions and to do work. As discussed in an earlier chapter, fatty acids are highly reduced so their oxidation by dioxygen is highly favored both enthalpically (exothermic reaction) and entropically. • 17.3: Ketone Bodies Thumbnail: Idealized representation of a molecule of a typical triglyceride, the main type of fat. Note the three fatty acid chains attached to the central glycerol portion of the molecule. (Public Domain; Benjah-bmm27 via Wikipedia) 17: Fatty Acid Catabolism Search Fundamentals of Biochemistry Introduction In this chapter, we will discuss the breakdown of fats to produce ATP. Most of the available chemical energy stored in fats is in the form of highly reduced fatty acids. One form of fatty acid-containing lipids comes from our diet, which includes triacylglycerols (TAGs) and membrane lipids. Fatty acids, mostly in the form of TAGs, are moved in the circulation in the form of large lipid-carrying vesicles called lipoproteins. The lipids can be imported into cells for storage and energy use. Another source of fatty acids comes from those synthesized within cells from the small molecule acetyl-CoA. Fatty acids are synthesized by an enzyme complex called fatty acid synthase. This enzyme is found most prevalently in adipose (fat) tissue and the liver. In addition, it is significantly expressed in the brain, lungs and mammary gland. TAGs, stored in lipid droplets, are found in most cells. The major tissue used for TAG storage is adipose (fat) tissue, whose volume consists mostly of lipid droplet(s). Given the large mass of muscle tissue, there is also a considerable amount of TAGs stored as small lipid droplets in muscle cells. However, skeletal muscle cells don't synthesize fatty acids. They have the genes for fatty acid synthase but do not transcribe it into RNA so no enzyme is made. They can however import them for catabolism. Muscle TAGs can be oxidized for energy, especially during endurance excercise. TAGs are also stored in the liver in lipid droplets. The liver also assembles lipoproteins, which are released by the liver. Excess TAGs are stored in the liver in various diseases including alcoholism and also in nonalcoholic fatty liver disease (NAFLD), which can progress into nonalcoholic steatohepatitis (NASH), a much worse disease. White and Brown Adipose Tissues There are two major forms of triacylglycerol-storing fat tissues, white adipose tissue (WAT) and brown adipose tissue (BAT). The more abundant WAT store triacylglycerols in one large lipid droplet in the cell and release fatty acid in processes controlled by the hormones insulin and epinephrine. This simple role can mask the fact that adipose tissue is a major player in the endocrine system and is involved in cell signaling and systematic control of metabolism. Adipose tissue releases the key hormones leptin and adipisin, which in analogy to the hormones and signaling agents released by immune cells (cytokines, lymphokines), can be called adipokines. They also secrete other adipokines including tumor necrosis factor α (TNF-α), adiponectin, and resistin. In contrast, BAT is specialized not to store and release fatty acids. but rather to oxidize fatty acids in ways that maximize heat production, preventing hypothermia. They have multiple smaller lipid droplets, displaying a larger surface area for lipolysis, the hydrolytic cleavage of fatty acids from the TAGs. A particular mitochondrial protein, uncoupling protein 1 (UCP1), is expressed in brown but not white adipocytes, allowing a "futile" metabolic cycle leading to dissipation of heat instead of ATP synthesis. The relative abundance of white and brown adipocytes is critical in diseases like obesity and type 2 diabetes. BAT tissue is especially important in small animals (and in newborns) for thermoregulation. For smaller organisms, the surface area to volume ratio is greater than the ratio for larger animals, allowing more heat loss. The ratio of surface area AS per volume V for a sphere is given by: $\dfrac{\mathrm{SA}_{\text {sphere }}}{\mathrm{V}_{\text {sphere }}}=\dfrac{4 \pi \mathrm{r}^{2}}{\left(\dfrac{4}{3}\right) \pi \mathrm{r}^{3}} \nonumber$ Let's assume an average large adipocyte is a sphere of diameter 100 uM. Compare this to a large sphere with a 100 times greater diameter (10,000 uM). The smaller sphere has a 1/100 of the diameter but a surface area/volume ratio 100 times greater than the large sphere. An intermediate type of fat tissue consists of "bright" adipocytes. White adipocytes can be coaxed to differentiate into bright and brown cells, which could be an obesity treatment. In this chapter section, we will follow the fate of fatty acids from dietary lipids which are cleaved from TAGs, loaded into chylomicrons, a lipoprotein assembled in the small intestine, secreted into the circulation, and taken up by the liver. The liver can store the incoming fatty acids in TAGs or release them back into the circulation in the form of another lipoprotein, very low-density lipoproteins (VLDL). Circulating VLDL can exchange lipids with other circulating lipoproteins. Lipoproteins deliver fatty acids to cells after interaction with the cell surface of target cells and either cleavage of TAGs by cell membrane-associated enzyme lipase, followed by fatty acid uptake, or by endocytosis of lipoproteins into the cells. Lipoproteins Before we look in more detail at the individual steps in lipids processing, let's look at the different lipoproteins, the large vesicular structures that allow the transport of fats, very insoluble molecules, in the circulation. Unlike normal liposomes or vesicles that have a lipid bilayer surrounding an interior aqueous compartment, lipoproteins have only a single monolayer of phospholipids encapsulating a nonaqueous interior filled with TAGs, cholesterol, and cholesterol esters. The protein part of the lipoprotein consists of one or several proteins bound on the outside of the particle. The proteins help solubilize the lipoprotein, confine its size, and prevent aggregation of the lipoproteins, which would be a health risk. The structure of a typical lipoprotein is shown in Figure $1$. Lipoproteins are classified based on density. The lowest density chylomicrons are the largest with the most lipids (mostly TAGs) in their interior compartment. Very large density lipoproteins (VLDL), intermediate density (IDL), low density (LDL) and high density (HDL) have decreasing size, less encapsulated lipids, and increasing density. The relative sizes are shown in Figure $2$. Lipoproteins (except chylomicrons) could be classified as nanoparticles, which typically vary in size from 1-100 nm. Larger lipoproteins as well as chylomicrons form emulsions in the blood, much as milk (also cloudy) is an emulsion of lipid/protein particles. The serum of people with high levels of lipids (hyperlipidemia) can look milky white, especially after eating foods rich in TAGs, when levels of chylomicrons are very high. Figure $3$ shows the blood of a patient with hyperlipidemia after the addition of EDTA (which binds Ca2+ and prevents clotting) that has settled (without centrifugation). The milky white plasma on top (lower density) most likely has high concentrations of chylomicrons and/or LDL. The lower layer contains mostly red blood cells. No x-ray structures of lipoproteins are available. However, a structure of a nascent HDL particle (3k2s) has been determined by small-angle neutron scattering. Figure $4$ shows an interactive iCn3D model of it. The major protein in HDL, a lipoprotein that protects against cardiovascular disease, is apolipoprotein A-I (apoA-I). Figure $4$ shows that it adopts an antiparallel double superhelix as it wraps around the nascent HDL. The more hydrophobic surfaces of apo A-I are oriented inward allowing interactions with hydrophobic lipids in the core. It is probably prototypical for nascent lipoproteins. It will give you an idea of how proteins wrap around the outside of the particle. Mature lipoproteins are most likely spherical. This nascent HDL in the model contains 200 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholines (POPC) molecules, 20 cholesterol, and a single copy of apolipoprotein A-I (apoA-I). Figure $5$ below shows a cartoon image of VLDL, assembled from lipids synthesized/taken up by and released from the liver, and chylomicrons, assembled from dietary fats and released from enterocytes in the small intestine (size of the lipoproteins is not to scale). Note that VLDL has one copy of Apo B-100 while chylomicrons have one copy of Apo B-48. All lipoproteins, except HDL, are members of the Beta-lipoprotein family as they contain an apo-B protein. The liver synthesizes apo B100, which becomes a permanent part of VLDL (i.e it is not exchangeable with other lipoproteins) and its metabolic derivatives so any lipoprotein containing apo B100 arose from the liver. Other proteins on lipoproteins are exchangeable. In contrast, enterocytes in the small intestine produce apo B-48 (48% the size of apo B100) so this protein marks the lipoproteins (chylomicrons and chylomicron remnants) that were assembled in the small intestine. Apo B100 has over 4500 amino acids and a molecular mass of 555K. The gene for the intestinal apo B48 is the same as for apo B100 except that it has a premature stop codon which leads to the shorter truncated apo B-48. The apoproteins bind to specific receptors on cells which may allow the uptake of the lipoprotein. For example, the LDL receptors bind to the apo B-100 protein on a region removed from the apo B48 protein of chylomicrons. It also binds ApoE, which is found mostly predominately on HDL and VLDL but some are present in LDL. The LDL receptors have also been called the ApoB/ApoE receptor. Over 90% of the apoB-containing particles in circulation are LDL. In addition, chylomicrons are present in circulation only after eating. Some apoproteins can act as cofactors and inhibitors for lipoprotein processing. Lipoproteins and Cardiovascular Risk High concentrations of LDL are associated with increased cardiovascular risk. Chylomicron levels, given their transient and lower concentration levels, do not pose a health risk unless the enzyme required to remove fatty acids from them, lipoprotein lipase, is missing or defective, or if another apoprotein component, apo CII, which mediates the interaction with lipoprotein lipase, is missing. LDL-C (a term used to describe the total cholesterol in LDL particles which is routinely measured in clinical labs) can be lowed by a healthy diet centered around plant food). Drugs like statins, which decrease endogenous cholesterol synthesis, also remarkably lower LDL levels and decrease cardiovascular risk. However, another protein, lipoprotein, also called Lp(a) or LP little a is an independent cardiovascular risk factor. Its blood concentration is regulated by genetics and not by diet. These particles contain, in addition to apo B100, apo (a), a protein that has a very unique repeating structure (up to 40 times) called a kringle, which is also found in some proteins involved in the blood coagulation system. People whose genes encode apo (a) with the fewest number of kringles express lots of that protein and their Lp(a) particles are smaller. This confers a greater cardiovascular risk compared to those expressing proteins with a large number of kringle domains. Figure $6$ (from Amgen) shows models of Lp(a) with different numbers of repeating kringle (kinked) domains. From a biochemical perspective, it is interesting to explore the differences in apolipoprotein binding to a single-leaflet encapsulated lipid nanoparticles compared to the interaction of peripheral and integral membrane proteins with intact bilayers (which we studied in Chapter 12.1). As mentioned above, the more nonpolar surfaces of apo AI in HDL are oriented inward toward the nonpolar lipid core. Presumably, apo B proteins in chylomicrons and LDL also wrap around the entire lipid surface. The major organizing scaffolding protein of HDL is apoA-I (see iCn3D model above), It presumably plays a role similar to apoB in chylomicrons and LDL, but it is exchangeable. (Note: ApoA-I is also found in chylomicrons.) It is also a cofactor for the enzyme lecithin:cholesterol acyl transferase (LCAT), which effectively converts free cholesterol in the single bilayer into esterified cholesterol esters within HDL. In its apo-form, it also interacts with the cell surface transporter ATP-binding cassette A1 (ABCA1), which plays a role in the assembly of HDL particles. HDL also has apo C and apo E proteins, all of which are exchangeable. It must be difficult to determine the structure of lipoproteins given their heterogeneity and size.  The apoproteins have hydrophobic surfaces that promote self-association and aggregation. Apolipoproteins in the A, C, and E classes have repeating amphiphilic helices which imbed to some degree in the lipid particles. In addition, the proteins have a significant disorder and can adopt many bound conformations. Figure $7$ shows an interactive iCn3D model of the AlphaFold predicted structure of human ApoA (P06727) The blue cartoon color represents high certainty in the AlphFold predicted structure while yellow to orange represents low certainty.  The hydrophobic side chains are shown as sticks and help illustrate the amphiphilic nature of the structure. Figure $8$ shows an interactive iCn3D model of the AlphaFold predicted structure of human ApoE (P02649) The blue cartoon color represents high certainty in the AlphFold predicted structure while yellow to orange represents low certainty.  The hydrophobic side chains are shown as sticks and help illustrate the amphiphilic nature of the structure. The exchangeable apolipoproteins have similar genetic sequences (four exons and three introns), as well as similar amino acid sequences. They have 11-mer amino acid tandem repeats and some (A-I, A-IV) have 22-mer tandem repeats. These repeats form amphiphilic helices as determined by sequence analysis. The first amino acid in the amphiphilic helix is often positively charged and a negative one is often found in the middle. Proline, a helix breaker, is often, but not always found between the helices. Figure $9$s shows the primary sequence of apoA-I. An 11-mer repeat is shown in yellow highlight. The other highlighted stretches (different colors) are 22-mer repeats. Note that the repeats are not of identical sequences but rather of sequences that can form amphiphilic helices (i.e. secondary structure repeats). The bottom part of Figure $9$shows a helical wheel projection (using Heliquest) of the red-highlighted 22-mer repeat. The arrow shows the hydrophobic moment with the arrowhead pointing to the more nonpolar face. The particular amphiphilic helix shown may or may not facilitate the binding of the bound conformation of the protein. It follows that the relative areas of the hydrophilic and hydrophobic faces in the amphipathic helixes influence the lipid-associating properties of the exchangeable apolipoproteins. Another factor that might influence the lipid-binding ability of exchangeable apolipoproteins and which has not been studied in detail so far is the arrangement of tandem repeating amphipathic helixes with respect to one another. Actual amphiphilic helices would bind to the membrane in a parallel fashion with the nonpolar face anchoring the protein to the lipid surface. Other experimental techniques are used to determine how a peptide or protein that can form amphiphilic helices interact with the lipid surface. These include site-directed mutagenesis studies coupled with spectroscopic (CD, fluorescence) and binding assay methods (using liposomes). The properties of a membrane-bound amphiphilic helix are affected by the exact size and distribution of the polar/charged and nonpolar side chains. On binding, they sense or cause membrane curvature, interact with specific lipids, and stabilize specific membrane conformations (such as spherical for lipoproteins). Figure $10$ shows how different proteins with amphiphilic membranes interact with membrane surfaces. A key point to note is the large conformational changes that occur as the protein or parts of it go from the free, more disordered state, to the bound state with lipid-associating amphiphilic helices. The following proteins are depicted in the figure. 1. The peroxisomal membrane protein Pex11 amphiphilic helix distorts the membrane; 2. ARF1 is a small G protein in which only the GTP form localizes and binds through an amphiphilic helix to the membrane; 3. The ALPS motif of the golgin GMAP-210 binds to only highly curved vesicles; 4. The yeast transcriptional repressor Opi1 binds to the endoplasmic reticulum (ER) membrane in part through an amphiphilic helix; 5. The heat shock protein Hsp12 has a long amphiphilic helix which helps stabilize the membrane; 6. The extremely long amphiphilic helix of perilipin 4 coats lipid droplets and stabilizes even if there is a lack of phospholipids. Dietary uptake and release into the circulation Now how are the lipid nanoparticles assembled? We'll start with dietary lipids in the form of TAGs, glycerophospholipids, and cholesterol esters. The figure below shows key steps which are described in Figure $11$ Here are some key steps depicted in the figure: • hydrolysis (lipolysis) of TAGs by pancreatic lipase, cholesterol esters (CE) by cholesterol esterase, and glycerophospholipids (GP)by phospholipase A2 in the lumen of the intestine. These enzymes interact at the interface of the lipid substrates and aqueous surroundings; • the resulting products, which include free fatty acids (FA), 2-monoacylglycerol (MG), free cholesterol (FC), and lyso-glycerolphospholipids (lyso-GP), aggregate with the help of bile salts to form emulsions (like oil drops in water), which can be taken up by diffusion or possibly endocytosis when present in high amounts. Alternatively, membrane transporters (like FABPs and other proteins) can move them into the cell by facilitated diffusion; • cytoplasmic transporters like fatty acid binding proteins move the lipolysis product to the ER where free fatty acids are reesterified. The enzymes involved include mono- and diacylglycerol acyltransferases (MGAT, DGAT) and sterol O-acyltransferase 2, also known as acyl-coenzyme A:cholesterol acyltransferase (ACAT-2). Multiple enzymes are involved in the resynthesis of glycerophospholipids); • Apo B-48 is synthesized by ribosomes bound to the ER and interacts with a heterodimer of microsomal triglyceride transfer protein large subunit two (MTP) and protein disulfide isomerase (PDI). This facilitates the folding of apo B48 and loading of lipids using MTP into pre-chylomicrons; • pre-chylomicron vesicles move to the Golgi with the help of Sar1b, a small G-protein (and GTPase) where the particle assembles to the full chylomicron, which is released from the cells as the mature large lipid nanoparticle. An intriguing feature of lipases is that they work at the interface between the aqueous and nonaqueous (in this case lipid nanoparticle) environments. Let's briefly consider the mechanism of hydrolysis of TAGs by equine pancreatic mechanism. This enzyme utilizes the same mechanism we have seen earlier for the hydrolysis of a peptide bond by serine proteases. A catalytic triad of Asp 176, His 263 and Ser 152 as a nucleophilic catalyst is shown in the partial reaction displayed in Figure $12$. An acyl-Ser intermediate forms in step B (above), after the collapse of an oxyanion intermediate in step A, to form the product diacylglycerol. In the second half of the reaction (not shown completely), water, in a hydrolysis reaction, cleaves the acyl-Ser intermediate to reform the active enzyme as it releases the free fatty acid, R3CO2H. Other lipases also employ the same catalytic triad. Figure $13$ shows an expanded diagram showing the flow and fate of lipoproteins. Chylomicrons interact with lipoprotein lipase (LPL), which also uses an Asp-His-Ser catalytic triad, to cleave fatty acid esters, which allows the delivery of free fatty acids to adipose cells. The adipocytes can also undergo de novo fatty acid synthesis. Fatty acids (FA) can also be produced by lipase-mediated lipolysis of stored TGs. Any of these free fatty acids (FA) in the adipocyte has two fates. They can be reesterified to glycerol to form TAGs (TG in the figure) or be exported from the cell and bind to a plasma carrier protein and transported to the liver, where it can be taken up by a variety of membrane proteins importers shown in the figure in yellow boxes. There, as in adipose cells, they can be reesterified to form TAG stores, which can then be packaged into VLDL particles for export. The fatty acid delivered (or synthesized) could also be used for ATP production through the citric acid cycle and oxidative phosphorylation. VLDL in circulation can undergo lipolysis by lipoprotein lipase to produce fatty acids for uptake in "extrahepatic" tissue (bottom right of the diagram). As fats are removed from VLDL, their density increases as it forms IDL and LDL, which could be considered VLDL "remnants". VLDL is very enriched in TAGs, but after metabolic processing, the resulting LDL is depleted in TAGs and enriched in cholesterol/cholesterol esters. LDL (not shown in the above figure) can be taken up (endocytosed) by the liver and other cells after binding to LDL receptors, which recognize apo-B100 and other apoproteins. This allows the delivery of predominately cholesterol and cholesterol esters to tissues. How do adipocytes and hepatocytes determine if free fatty acids should be esterifed for storage or released for energy use by other tissue? We'll discuss that in a subsequent section but the short answer is that in healthy fasting and exercise states, hormones (glucagon, epinephrine) will activate lipolysis in the liver and adipose cells, while in the fed state, insulin will promote storage of fatty acids as triacylglycerols. Adipose cells don't assemble and release lipoproteins. Instead they release free fatty acids in the circulation which are carried by albumin, the major serum/plasma protein in the blood. The iCn3D Figure $14$ shows an interactive iCn3D model of the complex of human serum albumin (HSA) binding seven 20:4Δ5,8,11,14 - arachidonic acids (1gnj ). Given the multiple binding sites for fatty acids in albumin, it should come as no surprise that albumin also binds a host of small drugs, including medicinal drugs and toxins such as warfarin (blood thinner), diazepam, ibuprofen, indomethacin, and amantadine.  These appear to bind preferentially at two major drug binding sites. This binding is probably helpful in delivering drugs through the circulation but potentially not useful if they aren't delivered to appropriate target tissue. We discussed the structure of micelles which are spherical assemblies of single-chain amphiphiles that act as detergents. Oil from your clothes can enter the nonpolar interior of the detergent micelle and effectively solubilize the nonpolar molecule in the micelle, which are effectively nanoparticles with a diameter of 5-15 nm. You should hence not be surprised to discover that lipoproteins can also carry fat-soluble vitamins, steroid-like endocrine-disrupting substances, and drugs. Lipoprotein lipase The enzyme that breaks now TAGs in circulating chylomicrons and VLDL is lipoprotein lipase (LPL). It is a soluble protein secreted by adipocytes and muscle cells but is made by many cell types. It works at the luminal side of blood vessel endothelial cells and is recruited to that membrane surface by binding to the glycosylphosphatidylinositol-anchored high-density lipoprotein-binding protein 1 (GPIHBP1) as well as the proteoglycan heparan sulfate at the cell surface. What is so interesting is that GPIHBP1 is only synthesized by endothelial cells. When lipoprotein lipase is secreted from cells, it binds to the extracellular matrix heparan sulfate but dissociates on the cleavage of heparan sulfate by heparinases. GPIHBP1 is highly acidic with an intrinsically disordered N-terminal domain containing a sulfated tyrosine and is highly enriched in glutamates and aspartates, which are often sequential in the sequence. Here is the single-letter sequence for amino acids 25-50 of the human version of GPIHBP1: EEEEEDEDHGPDDYDEEDEDEVEEEE. This sequence would have similar electrostatic and binding properties to the highly negatively charged heparan sulfate to which it also binds. LPL also binds Ca2+ which stabilizes the active dimeric form of the protein. Its enzymatic activity is activated by apoC-II. Like pancreatic lipase, it employs a Ser-132, Asp-156, and His-241 triad in its hydrolytic action on TAGs/ Figure $15$l below shows an interactive iCn3D model of LPL in complex with GPIHBP1, shown in brown (6E7K). The calcium ion is shown (grey spacefill) as well as the catalytic triad (labeled, sticks, CPK colors). The highly negatively charged stretch of amino acids in GPIHBP1 was not present in the crystal structure. LDL: Receptor and Uptake Lipoproteins are taken up into cells through receptor-mediated processes. Let's focus generically on the LDL receptor, the major carrier of cholesterol, given its role in cardiovascular disease. It is found in the cell membranes in most tissues. It is has many domain repeats, as illustrated in the Figure $16$ calculated by SMART. They include the N-terminal region cysteine-rich LDLa domains, which bind LDL, epidermal growth factor domains, LY (or LDLb) domains, and a transmembrane domain (blue rectangle). Figure $17$s shows an interactive iCn3D model of the extracellular domain of the LDL receptor (1n7d). Four tandem LY (LDLb) domains are shown in cyan, LDLa domains are shown in magenta and the EGF domain is shown in dark orange. Glycans are shown in symbolic nomenclature for glycans. Zoom into the structure to see the two disulfide bonds in each LDLa domain as well as the Ca2+ ions that stabilize the domains. LDL binds its receptor at a broad binding interface with multiple LDLa domains. This may account for the fact the lipid nanoparticles with apo B100 or apo E can bind to it. The binding triggers a series of signaling events that lead to internalization by endocytosis of the receptor in pits coated with the protein clathrin. These eventually fuse with lysosomes where they are degraded and cholesterol delivered to the cell. The steps are described in Figure $18$. The LDL receptor survives lysosomal degradation and along with newly made receptors is delivered to the plasma membrane continually. A key protein, proprotein convertase subtilisin kexin type 9 (PCSK9), a serine protease secreted by the liver, promotes enzymatic degradation of the receptor and prevents its recycling to the membrane. It also binds to VLDLR and apolipoprotein E receptors and promotes their degradation as well. Its action reduces LDL clearance from the blood, increasing cardiovascular risk, so inhibitors of its action might be potent drugs to decrease circulating LDL. The LDL receptor is just one member of the LDL receptor family. Other members of the family are illustrated in Figure $19$. These include the LDL receptor (abbreviated Ldlr in Figure $17$, as well as the VLDL receptor (Vldlr), apolipoprotein E2 receptor (Apoe2), and LDL receptor-related proteins (Lrp)1-4. These also have a NPxY-motif (asterisk in the cytoplasmic domain) and a YWTD/β-propeller domain. Given the similarity in domain structure for the LDL family of receptors, the conformational flexibility of the apolipoproteins (at least free in solution), and similar structures for the exchangeable apolipoproteins, it shouldn't be surprising that the LDL receptor would interact with different classes of lipoproteins, albeit with different affinities. As mentioned previously, apoE is found most abundantly on HDL and VLDL/chylomicrons and their remnants. It serves as a ligand that binds to members of the LDL receptors family (remember that LDL generally binds the LDL receptor through apo B100). Apo E and Alzheimer Apolipoprotein E has three major variants (alleles) named ε2, ε3, and ε4 (also called ApoE2, 3, and 4). ApoE3 is the most prevalent. ApoE4 is found in only 15% of people but more than 50% with Alzheimer's Disease (AD), so it's a risk factor for this disease. AD affects the brain, which also contains up to 30% of the cholesterol in the body, so aberrations to cholesterol transport and uptake in the brain are not unexpected in neurodegenerative diseases like AD. ApoE is secreted by brain microglia (immune) cells and astrocytes (specialized glial cells). It assembles lipids into lipoproteins (HDL-like) and becomes the major vehicle for binding to and importing into neurons in a process initiated by the apoE receptor. The major apoE receptor for clearance of lipoproteins in the brain is sortilin (SorLA in Figure $17$). AD is characterized by the accumulation of a toxic amyloid prion protein called amyloid beta (Aβ). It is derived from selective but abnormal proteolysis of the neural integral membrane protein amyloid precursor protein (APP). Aβ aggregates to form insoluble neurotoxic extracellular Aβ amyloid plaques. The process in normal and diseased cells is shown in Figure $20$. The figure shows normal (left) and aberrant processing of APP and the family of proteases (secretases) involved. While the LDLR doesn't appear to bind to APP or influence its proteolytic processing, it does bind Aβ. LRP1 is much bigger than LDLR, binds a multitude of ligands, and can be cleaved with the same enzymes as APP. Its expressed in the liver and especially in the brain and can regulate the removal of Aβ. Immune cells in the brain, called microglial, remove Aβ plaques (which are extracellular) by phagocytosis. ApoE4 increases the inflammatory response (as measured by cytokine release) of the microglia (a good thing if the responses prevents infection or rids Aβ plaques) but also inhibits their ability to phagocytose the Aβ plaques and their metabolic activity. An additional note:  Having one allele ApoE4 allele appears to increase the risk of severe COVID-19 five times while being homozygous for E4 leads to a 17-fold increased risk of severe disease. Scavenger Receptors Patients with homozygous familial hypercholesterolemia (FH) have very high levels of LDL derived from defects in binding and update. Patients display fatty acids streaks under vessel endothelial cells which morph into calcified plaques and lesions filled with fat. Monocytes/macrophages, which migrate to sites of vascular injury, take up LDL and eventually differentiate into foam cells filled with lipids. Somehow, they have receptors that can bind and internalize LDL when the "normal" LDL receptor can't. Brown and Goldstein found that a specific chemical modification of LDL, acetylation, was necessary for the rapid uptake of "modified" LDL into macrophage receptors. These receptors are now called scavenger receptors (SRs). There is a large family of scavenger receptors. It consists of classes A-J proteins that share functional but not sequence homology. They are found on macrophages and endothelium. They bind to and help remove "damage" signals including damage-associated molecular patterns (DAMPs) and chemical species chemically modified by reactive oxygen species. The ligands are often polyanions, end-stage glycans, and extracellular matrix proteins. One such example is oxidized-LDL (produced in vivo or by chemical oxidative modification with malondialdehyde), which binds to the same scavenger receptor, SR-A1, also called Macrophage scavenger receptor type I, as acetylated-LDL. Figure $21$ shows the domain structure of the SR family. SR-A1/MSR1 not only binds acetylated and oxidized LDL but also β-amyloid (), heat shock proteins (), and PAMPs from some bacteria and viruses. It's very difficult, given the ever-increasing amount of "omic" data (genomic, proteomic, lipidomics, interactomics, metabolomics), for readers and authors alike, to conceptualize all of the possible combinations of interactions among biological molecules. For visual learners and perhaps everyone else, it's extremely useful to portray information on structures and interactions visually. An example using STRING, a database of known and predicted protein interactions, for the domain structure and protein:protein interactions of SR-A1/MSR1, is shown in Figure $22$. Note the interactions with apolipoproteins, apoB, apoE, and apoA1. The right hand side of the figure also shows interactions with collagen alpha-2(IV) chain (COL4A2), which is found in the extracellular matrix. HDL metabolism: The Good Cholesterol High levels of LDL (and Lp(a)) pose a cardiovascular risk. In contrast, high levels of HDL and apo A-I are cardioprotective. HDL is involved in "reverse" cholesterol transport as it is taken up by the liver and sent to the intestines for elimination from the body. We have shown earlier in Figure 2 that HDL exists as many variants, reflecting the assembly and remodeling of HDL by enzymes and lipid transfer proteins. Figure $23$ shows the lifecycle of HDL. Secreted apo A-1 accretes lipids in the circulation through the transport and delivery of phospholipids and cholesterol from cell membranes by the ATP-binding cassette transporters (ABC) A1 and G1. Another protein, the scavenger receptor BI (SR-BI), a polytopic integral membrane protein, is also involved. It acts as a receptor for a variety of "lipid" ligands including phospholipids, cholesterol esters, and phosphatidylserine (an outer membrane marker for cell apoptosis) as well as lipoproteins such as HDL. Other proteins are involved as well in both the assembly but especially in the remodeling of HDL. The lipolytic enzyme lecithin-cholesterol acyltransferase (LCAT) removes a fatty acid from phospholipids and adds it to free cholesterol in the HDL to form cholesterol esters. Two major lipid transfer proteins, phospholipid transfer protein (PLTP) and cholesterol ester transfer protein (CETP), move lipids between HDLs and other lipoproteins. Cholesterol ester transport protein is made and secreted from the liver. It appears to exchange cholesterol esters from HDL for the return of TAGs from VLDL. Other enzymes (lipoprotein lipase and hepatic lipase are also involved in forming free fatty acids. In the final step, HDL can deliver cholesterol ester and cholesterol to the liver through binding to liver scavenger receptor BI (SR-BI) mediated by apo A-I. The protein is expressed by the liver, adrenal gland, endothelial cells, macrophages, and many other tissues. It appears that HDL is not primarily taken up by the liver by endocytosis. In contrast, LDL is taken up by endocytosis mediated by the LDL receptor. SR-BI facilitates the transfer of cholesterol esters from bound HDL to the liver cell. HDL and cardiovascular risk Unlike the widespread use of statins, which reduce LDL-C concentrations and clinically reduce cardiovascular disease risk, drugs (fibrates, niacin, inhibitors of cholesterol ester transfer protein - CETP), which raise HDL levels, don't seem to lead to significant decreases in cardiovascular risk. The cholesterol delivered in excess to macrophages can lead to the formation of foam cells under the endothelial layer.  Foam cells are proinflammatory and convert in time to cholesterol plaques. In contrast, HDL-C does appear to have this direct atherogenic effect.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/17%3A_Fatty_Acid_Catabolism/17.01%3A_Digestion_Mobilization_and_Transport_of_Fats.txt

Search Fundamentals of Biochemistry Introduction Fatty acids, esterified to glycerol in triacylglycerols, are the major source of stored energy in organisms. As we burn fossil fuels to produce energy to drive our society, so can we "burn" fatty acids to indirectly produce for heat, ATP to drive biosynthetic reactions, and to do work. As discussed in an earlier chapter, fatty acids are highly reduced so their oxidation by dioxygen is highly favored both enthalpically (exothermic reaction) and entropically. Of course, the biological oxidation reactions occur in a stepwise fashion using not O2 directly, but rather less potent oxidizing agents like NAD+ and FAD. We'll focus first on fatty acid oxidation in animals (humans). As we discussed in the previous section, fatty acids released from triglyceride stores on signaling by epinephrine and glucagon in exercise and between meals are used for energy when glycogen stores are low and without breaking down protein in muscles to produce energy. Some fatty acids are broken in the normal process of membrane turnover and removal of xenobiotic lipids. Most fatty acids are oxidized in the mitochondria, where the oxidation reaction occurs at the beta-carbon of the acyl chain, as shown in Figure $1$. This pathway is called β-oxidation. Fatty acids are oxidized in a step-wise fashion by this pathway. In each repetitive cycle of this pathway, acetyl-CoA and one CO2 are released. In addition, oxidation can occur at both the alpha- and beta carbons when oxidized in an organelle called the peroxisome. The α-oxidation pathway is used for fatty acid branched at the beta carbon 3 releasing one CO2 until the beta-oxidation pathway can be used. The peroxisome degrades fatty acids that can't be oxidized in the mitochondria. These include very long-chain fatty acids (VLCFAs) like 24:0 and 26:0, and in addition, branched-chain fatty acids (BRCHAs) including some fatty acids from dietary sources such as pristanic acid (an odd-chain 15:0 fatty acid methylated at carbons 2, 6, 10, and 14). The α-oxidation pathway can't be used to completely oxidized fatty acids in the peroxisome. At some point in the oxidative stepwise pathway, the resulting shorter fatty acids are exported to the mitochondria for β-oxidation. Also, enzymes in the endoplasmic reticulum have the ω-oxidization pathway which oxidizes fatty acids at the omega or terminal carbon. The enzyme used is the monooxygenase cytochrome P450 which uses one oxygen from O2 to hydroxylate the ω-carbon. Peroxisomes - An underappreciated organelle These organelles, initially called microbodies, are vital to cellular metabolism and health. In people with Zellweger syndrome spectrum, there is a severe disorder in the formation of peroxisomes, which is often lethal. They are important metabolically in lipid metabolism, synthesis of myelin sheath lipids, and metabolism of reactive oxygen species like peroxides. The enzymes catalase and urate oxidase are found in such high concentrations they often form crystal "bodies" in the matrix of the peroxisome. Additional roles include responses to pathogens and viruses. Effectively they are a protective organelle. In contrast to mitochondria, peroxisomes, like most other organelles, have a single bilayer and no DNA, from which transcription of RNA and translation of proteins occur. All proteins are hence imported from the cytoplasm after synthesis on free ribosomes. Imported proteins have a peroxisome targeting sequence (PTS) of serine-lysine and leucine (SKL) near their C-terminus which facilitates the binding of the proteins to a PTS receptor in the peroxisome membrane. These organelles oxidize very long-chain fatty acids (VLFA), make and break down hydrogen peroxide (hence the name), and also synthesize plasmalogens. The enzymes involved in the stepwise cycle of reactions in the peroxisome β-oxidation pathway use enzymes different from those used in the mitochondria for β-oxidation. For those more inclined towards chemistry than biology, yet another organelle with its structures and function may seem like one too many. However, this less-discussed organelle is critically important in its own right. Figure $2$ shows features of peroxisomes and their proteins. One interesting feature is its relationship with different organelles in cells, as shown in the left panel of Figure 2. Some proteins involved in organelle functions are shown as well (right panel). The peroxisome (PO, green) has binding interactions (red interfaces ) with the endoplasmic reticulum (ER) lysosomes, mitochondria, lipid droplets (a pseudo-organelle), and also itself (left figure). Some of the key membrane proteins (which we have discussed previously) involved in peroxisome function include the ABC transporter proteins ABCD1-3 for fatty acids transport, OCTN3 for organic and cation/carnitine transport, and MCT1/2 for monocarboxylate transport. In addition, peroxisomes have receptors for protein import mediated by PTSs and for peroxisome movement along microtubules in the cell. Mitochondrial β-Oxidation Mitochondrial β-oxidation in muscle generates acetyl-CoA, which enters the citric acid cycle for subsequent production of ATP through mitochondrial electron transport and oxidative phosphorylation. In the liver, the generated acetyl-CoA is used for ketone body production under fasting states. Figure $3$ shows the β-oxidation pathway for palmitic acid (16:0), a saturated fatty acid, starting with its import from the cytoplasm. The pathway involved cyclic removal of 2C unit until 16:0 is cleaved 7 times producing 8 2C acetyl-CoAs. The net chemical equation of beta-oxidation of 16:0 is shown in the equation below. \mathrm{C}_{16}-\mathrm{CoA}+7 \mathrm{NAD}^{+}+7 \mathrm{FAD}+7 \mathrm{CoASH}+7 \mathrm{H}_{2} \mathrm{O} \rightarrow 8 \mathrm{Acetyl}-\mathrm{CoA}+7 \mathrm{NADH}+7 \mathrm{FADH}_{2}+7 \mathrm{H}^{+} Figure $4$ shows an abbreviated comparison of the β-oxidation pathways in the mitochondria and peroxisomes. Peroxisomal beta-oxidation is used to metabolize very-long-chain fatty acids (VLCFAs), which are composed of 24-26 carbon units as well as branched-chain fatty acids (BRCHAs). It should be noted that a likely NAD+/NADH mitochondria transporter, a multi-pass inner mitochondrial membrane protein has just been identified The transporter, MCART1, is also called SLC25A51, β-Oxidation - Mechanisms Fatty acids are imported into the matrix from the cytoplasm through their acyl-CoA derivatives. Two different proteins are required for their import. One is carnitine palmitoyltransferase-1(CPT-1), which transfers the acyl group from CoASH to a carrier protein carnitine. The acylcarnitine is translocated through the inner membrane by the carrier protein carnitine-acylcarnitine translocase (CACT). Once inside the matrix, the acyl group is transferred back to CoASH by carnitine palmitoyltransferase-2 (CPT-2). This carnitine cycle is illustrated in Figure $5$. Figure $5$: Carnitine cycle and the connections between major catabolic pathways. At the outer mitochondrial membrane (OMM), fatty acyl-CoAs become linked to carnitine through carnitine palmitoyltransferase-1 (CPT-1). The complex is translocated across the inner mitochondrial membrane (IMM) via carnitine-acylcarnitine translocase (CACT). In the mitochondrial matrix, CPT-2 converts fatty acylcarnitines back to fatty acyl-CoAs, which enter the β-oxidation pathway. Free carnitine moves back into the cytoplasm through exchange with acyl-carnitines with CACT. β-oxidation in the matrix produces acetyl-CoA, which is also made from glycolytic pyruvate through pyruvate dehydrogenase. Hence acetyl-CoA links both glycolysis and fatty acid oxidation. The resulting acetyl-CoA can enter the TCA cycle when energy is needed. The mitochondrial carnitine/acylcarnitine carrier protein helps transport acylcarnitines of different lengths across the mitochondrial inner membrane for β-oxidation into the mitochondrial matrix. Figure $6$ shows an interactive iCn3D model of the human mitochondrial carnitine/acylcarnitine carrier protein AlphaFold model (O43772) Figure $6$: Mitochondrial carnitine/acylcarnitine carrier protein AlphaFold model (O43772). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...DdDNu9xVJGg2i9 The transmembrane helices are shown in gray. The N- (Met 1) and C-terminal (Leu 301) amino acids are shown in spacefill color CPK. Malonyl-CoA produced in the first committed step in fatty acids synthesis inhibits CPT1. This should make biological sense since fatty acid oxidation should not occur as fatty acids are synthesized. Palmitoyltransferase II (CPT II), which serves to convert acylcarnitine to fatty acyl CoA, traps the molecules within the mitochondrial matrix. In contrast to this regulated transport mechanism, very long chain fatty acids (VLCFAs) and branched-chain fatty acids are transported into peroxisomes by the ABCD1-3 transporters through an ATP-dependent process Mitochondrial β-oxidation of fatty acids has four steps that occur in the mitochondrial matrix. In those steps, a 16:0 fatty acid (for example) is converted to a (14):0 fatty acid and the 2C molecule acetyl-CoA. The (14):0 fatty acid undergoes 6 more rounds of the β-oxidation cycle until the entire 16:0 fatty acid is fully converted to 8 acetyl-CoAs. Step 1: Acyl-CoA dehydrogenase There are long- (LCAD), medium-(MCAD), and short-chain acyl-CoA dehydrogenases (SCAD) which catalyze the first oxidative step in the β-oxidation pathway. These enzymes catalyze the formation of a trans double bond between the α and β carbons (C2 and C3) on the acyl-CoA substrates. A stronger oxidizing agent than NAD+ is required to form an alkene between the two methylene groups, so FAD is used. Eventually the reduced FADH2 produced will lead to the production of 1.5 equivalents of ATP in the mitochondrial electron transport chain/oxidative phosphorylation. Figure $7$ shows the key oxidative step in the mechanism of acyl-CoA dehydrogenase to 2-enoyl-CoA by acyl-CoA dehydrogenases Figure $8$ shows an interactive iCn3D model of the medium-chain acyl-CoA dehydrogenase from pig liver mitochondria with octanoyl-CoA, a substrate (3MDE) Figure $8$: Medium-chain acyl-CoA dehydrogenase from pig liver mitochondria with octanoyl-CoA substrate (3MDE). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...SiAZExwXVV2TG6 Just two subunits of the biologically active tetramer are shown (dark gray and cyan). FAD is shown in each subunit (sticks, CPK colors, labeled). A bound substrate, octanoyl-CoA (spacefill, CPK colors, labeled CO8) is also shown in each subunit. The catalytic base, Glu 376, is shown in sticks, CPK colors, and labeled. The structures of the unliganded and acyl-CoA forms of the enzymes are very similar, so there are no large conformational changes on binding octanoyl-CoA. The ligand binds to the enzyme at the rectus (re) face of the FAD with the acyl chain buried. The fatty acyl chain of the thioester substrate is buried inside of the polypeptide and the 3'-AMP moiety is close to the surface of the tetrameric enzyme molecule. The carbonyl oxygen of octanoyl-CoA interacts with the ribityl 2'-hydroxyl group of the FAD and the main-chain carbonyl oxygen of Glu-376. Glu-376 acts as a general base as it removed the alpha proton in the reaction. Step 2. Enoyl CoA hydratase This enzyme catalyzes a hydration step of the double bond between the α and β carbons (C2 and C3), adding an OH group to the β carbon, in a reaction that poses little energy barrier. A likely mechanism for enoyl-CoA hydratase is shown in Figure $9$. Figure $10$ shows an interactive iCn3D model of the rat enoyl-CoA hydratase in complex with hexadienoyl-CoA (1MJ3) Figure $10$: Rat enoyl-CoA hydratase in complex with hexadienoyl-CoA (1MJ3). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...byx3eEKewEeEz9 Only one subunit of the biological hexamer is shown for clarity. Two glutamic acids (141 and 164) appear to activate a water molecule for the hydration reaction. Alanine 98 and Gly 141 appear also to be situated in an oxyanion hole which stabilizes the transition state and intermediate. The addition of the water is syn since the proton and OH group are added to the same size of the double bond. The glycine amide NH provides a strong hydrogen bond to the carbonyl of the substrate, hexadienoyl-CoA, helping to polarize the ene-one. The substrate trans-2-crotonyl-CoA is converted to the 3(S) alcohol instead of the 3(R) alcohol by a huge factor. Both the cis and trans isomers of a substrate analog (hexadienoyl-CoA) can bind to the enzyme, but only the cis isomer is polarized. Since the transition state is polarized as well, it would appear the bound cis isomer is strained and destabilized, suggesting that its binding is an example of transition state binding catalysis. Step 3. Beta-hydroxyl acyl CoA dehydrogenase After the addition of the OH on C3 (beta) OH during the hydration reaction, the resulting ROH is oxidized to a ketone, β-ketoacyl-CoA, by the oxidizing agent NAD+ using the enzyme β-hydroxyl acyl CoA dehydrogenase. The resulting NADH is reoxidized to NAD+ through the mitochondrial electron transport chain, which leads to the formation of 2.5 molecules of ATP for each NADH. Figure $11$ shows a plausible mechanism for the beta-hydroxyl acyl CoA dehydrogenase-catalyzed reaction. His 158 acts as a general base. Glu 170 increases the basicity of His 158. The other group is involved in H-bond and electronic stabilization interactions. Step 4. Acetyl-CoA acetyltransferase, mitochondrial - ACAT1 (also called 3-ketoacyl-CoA thiolase)_ The final step in the beta-oxidation pathway involves cleavage of the bond between the alpha and beta carbon by CoASH. This step is catalyzed by beta-keto thiolase and is a thiolytic (as opposed to cleavage by water - a hydrolysis) reaction. The reaction produces one molecule of acetyl CoA and a fatty acyl CoA that is two carbons shorter. The process repeats until the even chain fatty acid is completely converted into acetyl CoA. The activity of the enzyme is reversible and it can also catalyze the Claisen condensation of two acetyl-CoA molecules into acetoacetyl-CoA, as we will see in the synthesis of ketone bodies in the next chapter section. The reaction starts with the acylation reaction of the nucleophilic Cys 89 with the carbonyl at the 3-oxoacyl-CoA, with the concomitant release of acetyl-CoA. This forms a Cys 89-acyl covalent intermediate. In the next step, Cys 378 acts as a general base to facilitate the nucleophilic attack of free CoASH on the acyl-intermediate. His 348 acting as a general acid protonates the thiolate leaving group. The amino acids (Cys89, Cys378, and His348) are generally conserved in thiolases (Bhaskar et al. 2020). Kinetically this mechanism is a ping-pong reaction. A reaction mechanism is shown in Figure $12$. Figure $13$ shows an interactive iCn3D model of Human Mitochondrial 3-Ketoacyl-Coa Thiolase (T1) (4C2J) Figure $13$: Human Mitochondrial 3-Ketoacyl-Coa Thiolase (T1) (4C2J). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...dzAHKEY77WpNi7 Two subunits in the biological function dimer are shown (cyan and gray). The active site is shown in the gray subunit as CPK-colored stick and labeled. The numbers are a bit different than shown in the mechanistic figure. CoASH is shown in each subunit as sticks. The fatty-acyl tail appears to bind in a tunnel. A few more enzymes are needed Steps 1 through 4 outlined above apply to the beta-oxidation of a saturated fatty acid with an even-numbered carbon skeleton. Unsaturated fatty acids, such as oleate (18:1) and linoleate (18:2), contain cis double bonds that must be isomerized to the trans configuration by the enzyme enoyl CoA isomerase or reduced by the enzyme NADPH (2,4-dienoyl CoA reductase or 24DCR), using NADPH. Enoyl CoA isomerase This enzyme catalyzes the isomerization of cis double bounds to the trans form which mimics those formed by acyl-CoA dehydrogenase by FAD in step 1. Figure $14$ shows a plausible mechanism for the conversion of cis double bonds to their trans isomer. Glu 136 acts as a general base, while amide Hs of Leu 66 and Gly 111 stabilized the intermediate oxyanion and hence the developing charge in the transition state. They are optimally situated in the oxyanion hole. Figure $15$ shows an interactive iCn3D model of the Human mitochondrial Δ32-enoyl-CoA isomerase (1SG4) Figure $15$: Human mitochondrial Δ32-enoyl-CoA isomerase (1SG4) (Copyright; author via source). Click the image for a popup or use this external link:https://structure.ncbi.nlm.nih.gov/i...H6LxEzBKajq8b7 The three subunits are shown in different colors. The substrate analog octanoyl-CoA is shown in spacefill CPK colors bound to the gray subunit. The catalytic residues are shown in sticks with CPK colors and labeled. The distal omega end binds in a hydrophobic tunnel. 2,4-dienoyl CoA reductase or 24DCR An alternative way to deal with the cis double bond is simply to reduce it, in this case, NADPH. This enzyme is used on all C=C at the even-number position and more at odd-numbered positions. A mechanism for the reduction is shown in Figure $16$. Figure $17$ shows an interactive iCn3D model of a monomer of the homotetrameric human mitochondrial 2,4-dienoyl-Coa reductase (1W6U) Figure $17$: Monomer of the homotetrameric human Mitochondrial 2,4-Dienoyl-Coa Reductase (1W6U) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...pQueKc84sFqzt7 The model shows bound NADP+ and the substrate trans-2,trans-4-dienoyl-CoA. The active site is open enough to accommodate fatty acids of different lengths. Tyr-199 and Asn-148 stabilize the enolate and the oxidized nicotinamide. For odd-number chain fatty acids, propionyl-CoA to succinyl-CoA Although most fatty acids of biological origin have even numbers of carbons, not all of them do. Oxidation of fatty acids with odd numbers of carbons ultimately produces an intermediate with three carbons, propionyl-CoA, which cannot be oxidized further in the beta-oxidation pathway. These additional steps are necessary: 1. carboxylation to make (S)-methylmalonyl-CoA; 2. isomerization to (R)-methylmalonyl-CoA; 3. rearrangement to form succinyl-CoA. The last step of the process utilizes the enzyme methylmalonyl-CoA mutase, which uses the B12 coenzyme in its catalytic cycle. Succinyl-CoA can then be metabolized in the citric acid cycle. Figure $18$ shows the pathway for the conversion of propionyl-CoA to succinyl-CoA. Recent Updates:  7/24/23  Mechanisms for conversion of propionyl-CoA to succinyl-CoA. We will look at each of the enzymes in turn. Propionyl-CoA carboxylase The net reaction for this enzyme is shown in Figure $19$ below. Propionyl-CoA + ATP + HCO3- →  (S)-methylmalonyl-CoA + ADP + Pi + H+ Figure $19$: Propionyl-CoA carboxylase reaction The enzyme has 3 subunits.  The alpha subunits has biotin carboxyl carrier protein and biotin carboxylase domains.  The beta subunit has carboxytransferase activity.  The mechanism for the Streptomyces coelicolor propionyl-CoA carboxylase (PCC) is shown below in two parts. In Part 1, biotin is carboxylated by HCO3- to form an activated bicarbonate derivative (similar to an anhydride) that is high energy in comparison to its hydrolysis product, as shown in Figure $20$ below. This reaction takes place in the alpha subunit. Figure $20$: Formation of carboxylated biotin by propionyl-CoA carboxylase - Part 1.   M-CSA.  Gemma L. Holliday, Daniel E. Almonacid, Jonathan T. W. Ng.  Creative Commons Attribution 4.0 International (CC BY 4.0) License In Part 2, shown in  Figure $21$ below, the carboxyl group on biotin is transferred to propionyl-CoA to form methymalonyl-CoA. This reaction occurs in the beta subunit. Figure $21$: Part 2 - Formation of (S)-methylmalonyl-CoA by activated CO2 transfer from carboxybiotin by propionyl-CoA carboxylase. Holliday et al., ibid. The roles of the main chain atoms of Ala 182 and Gly 183 in the alpha subunit are to stabilize the propionyl-CoA while the backbone atoms of Gly 429 and Ala 430 in the beta subunit are to stabilize the oxyanion of carboxylated biotin. Figure $22$ shows an interactive iCn3D model of biotin and propionyl-CoA bound to Acyl-CoA Carboxylase Beta Subunit from S. coelicolor (1XNY) Figure $22$: Monomer of the biotin and propionyl-CoA bound to Acyl-CoA Carboxylase Beta Subunit from S. coelicolor (1XNY). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...K46r5EdZezF4o6 Biotin (labeled BTN) and propionyl-CoA are shown in spacefill in CPK colors. Methylmalonyl-CoA Epimerase The net reaction is shown in Figure $23$ below. Figure $23$: Methylmalonyl-CoA Epimerase reaction This reaction proceeds through an enolate intermediate after an abstraction of a proton from the chiral center of the methylmalonyl-CoA, as shown in the reaction mechanism for the enzyme from propionibacterium freudenreichii subsp. shermanii is shown Figure $24$ below. The reaction is readily reversible. Figure $24$:  Mechanism for methylmalonyl-CoA Epimerase.  Gemma L. Holliday et al. https://www.ebi.ac.uk/thornton-srv/m-csa/entry/33/.    Creative Commons Attribution 4.0 International (CC BY 4.0) License Note the presence of a Co2 ion in the active site. Figure $25$ shows an interactive iCn3D model of Methylmalonyl-CoA epimerase in complex with 2-nitronate-propionyl-CoA from S. coelicolor (6WFI) Figure $25$: Methylmalonyl-CoA epimerase in complex with 2-nitronate-propionyl-CoA from S. coelicolor (6WFI). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...Sqc98CCjAvzkp7 The enzyme is a dimer with two identical subunits and catalytic. One of the subunits is shown with a blue transparent surface with the bound 2-nitronate-propionyl-CoA inhibitor. The other is shown in gray cartoon with the key side chains involved in substrate binding and catalysis shown as color sticks and labeled. Methylmalonyl-CoA Mutase In this reaction, a methyl group is removed from  (R)-methylmalonyl-CoA to form succinyl-CoA, a shown in Figure $26$ below. Figure $26$: Methylmalonyl-CoA reaction This reaction going in the opposite direction is an example of a methyltransferase.  Another similar enzyme (which we will see in Chapter 18.4 - Amino Acid Degradation) catalyzes a methyl transfer from homocysteine to a new cofactor, cobalamin, which transfers it to cysteine to form methionine in a reaction catalyzed by methionine synthase.  Cobalamine and its methylated form are derivatives of Vitamin B12.  We will leave details of cobalamin biochemistry to the next chapter and will present a mechanism for methylmalonyl-CoA mutase here. We present the reaction for the reverse reaction, the conversion of succinyl-CoA to (R)-methylmalonyl-CoA by the enzyme from Propionibacterium freudenreichii subsp. shermanii in two parts below.  The enzyme is a heterodimer of an alpha and beta subunit and has a cofactor, adenosylcobalamin (coenzyme B12). In contrast, the human mutase, a homodimer, is very similar to the alpha subunit. First, a free radical is formed from the adenosyl group on the cofactor.  This promotes a free-radical rearrangement of succinyl-CoA to (R)-methylmalonyl-CoA (or the reverse for the pathway of interest here).  The reaction is shown in two parts for more optimal viewing. Figure $27$ below the first parts of the reaction for the conversion of succinyl-CoA to (R)-methylmalonyl-CoA by the mutase from Propionibacterium freudenreichii subsp. shermanii Figure $27$: Part 1 of conversion of succinyl-CoA to (R)-methylmalonyl-CoA by methylmalonyl-CoA mutase.  Gemma L. Holliday, Gail J. Bartlett, Daniel E.  Almonacid.  M-CSA.  https://www.ebi.ac.uk/thornton-srv/m-csa/entry/62/.  Creative Commons Attribution 4.0 International (CC BY 4.0) License Figure $28$ below shows the rest of the reaction for the conversion of succinyl-CoA to (R)-methylmalonyl-CoA. Figure $28$: Part 2 of the conversion of succinyl-CoA to (R)-methylmalonyl-CoA by methylmalonyl-CoA mutase. Gemma L. Holliday et al., ibid. The very last product from the bottom left reaction, the reformed active adenosylcobalamin cofactor, is not shown. The enzyme facilitates the hemolytic cleavage of the Co-C bond. This is followed by a free-radical rearrangement. Figure $29$ shows an interactive iCn3D model of the methylmalonyl-coenzyme A mutase from from Propionibacterium freudenreichii subsp. shermanii (1REQ) Figure $29$: Methylmalonyl-coenzyme A mutase from from Propionibacterium freudenreichii subsp. shermanii (1REQ). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...qPYm2sXjraCuN8 An analog of CoASH, desulfo-CoA (DCA) , is shown in spacefill, CPK colors and labeled.  Adenosylcobalamin is shown in colored sticks and labeled B12.  The alpha subunit is shown in gray with the key amino acids from the enzyme mechanism presented above shown as colored sticks and labeled.  The beta subunit is shown in blue. Propionyl-CoA is also produced as a product of the oxidation of methionine, valine, isoleucine, and threonine. (See the amino acid metabolism chapter for more details on mechanisms.) Very long chain oxidation In contrast to the oxidation of short and medium-chain fatty acids, which under beta-oxidation require four different discrete enzymes, the oxidation of very long-chain fatty acids (VLCFs) is carried out by two proteins with the second protein expressing three enzyme activities. The first step, analogous to step 1 in beta-oxidation described above, is carried out by a very long chain acyl-CoA dehydrogenase (VLCAD). The next three reactions are carried out by a single trifunctional protein (TFP) with two subunits. The α-subunit carries out the hydration (2-enoyl-CoA hydratase, ECH) and next oxidation step (3-hydroxyl-CoA dehydrogenase (HAD), while the β-subunit has 3-ketothiolase (KT) activity. Deficiencies of TFP can cause significant disease and even death. TFP is a heterotetramer of two α and two β subunits with the beta subunits forming a central homodimer. The two alpha units bind at each end and the whole complex forms an arc. There appears to be a "tunnel" allowing substrate transfer after each step. This minimizes premature intermediate release. Figure $30$ shows an interactive iCn3D model of the human mitochondrial trifunctional protein, a fatty acid beta-oxidation metabolon (6DV2) Figure $30$: Human mitochondrial trifunctional protein fatty acid beta-oxidation metabolon (6DV2). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...XiUXru4vkRVh87 • Grays: two thiolase subunits: reversible thiolytic cleavage of 3-ketoacyl-CoA into acyl-CoA and acetyl-CoA, a 2-step reaction involving a covalent intermediate formed with a catalytic cysteine. The catalytic residue (C138, C458, and H428) are shown as sticks with CPK colors. • Cyan and magenta subunits: enoyl-CoA hydratase (EC 4.2.1.17) and 3-hydroxyacyl-CoA dehydrogenase (EC 1.1.1.35) • The red dots represent the inner leaflet of the inner membrane so the proteins reside in the mitochondrial matrix. Regulation of beta-oxidation We saw that the metabolic decision to use carbohydrate energy reserves (glycogen) is a highly regulated event. Glycogen breakdown occurs during fasting and energy need. Fatty acids, our largest stores of energy, are released from triglyceride reserves in adipose cells through extracellular epinephrine and glucagon activation of pathways that activate intracellular hormone-sensitive lipase. Released fatty acids are bound to the serum protein albumin which transports them to tissue. Also as we mentioned above, malonyl-CoA inhibits the movement of fatty acids into the mitochondria. Malonyl-CoA is the first committed product of fatty acid biosynthesis. Each acyl-CoA product of each of the four enzymes engages in product inhibition for the enzyme that produced it. 3-ketoacyl-CoA also inhibits enoyl-CoA hydratase and acyl-CoA dehydrogenase [17]. The NADH/NAD+ and acetyl-CoA/CoA ratios also influence the beta oxidation pathway through allosteric regulation.   For example, the ratio of acetyl-CoA/CoA ratio affects the activity ketoacyl-CoA thiolase. Fatty acids also bind to the transcription factors called peroxisome proliferator-activated receptors (PPARs) and also coactivator PGC-1α, which regulate the transcription of enzymes in the beta-oxidation pathways. PPARs and the retinoid X receptor form heterodimers that bind to the PPAR response element in key promoters site involved in fatty acid degradation. These include CPT1, long-chain acyl-CoA dehydrogenase (LCAD), medium-chain acyl-CoA dehydrogenase (MCAD), and acyl-CoA synthetase (ACS). The PPARs have tissue specificity. We will discuss PPARs in more detail in the chapter on fatty acid synthesis. Peroxisomal α-Oxidation Alpha oxidation of fatty acids occurs in the peroxisome. It is used to metabolize phytanic acid (3,7,11,15-tetramethyl hexadecanoic acid), found in dairy products, animal fat, and some fish. Phytanic acid is produced in ruminants on the degradation of plant material and derives from phytol, an isoprenoid alcohol esterified to chlorophyll. Phytol is first converted to phytanic acid. Fatty acid β-oxidation can also occur in peroxisomes. In animals, peroxisomes are believed to be important in the initial breakdown of very long-chain fatty acids and methyl-branched fatty acids [11]. The enzymes involved in fatty acid oxidation in peroxisomes are different from mitochondria. An important difference is acyl-CoA oxidase, the first enzyme in peroxisome β-oxidation, which transfers the hydrogen to oxygen producing H2O2 instead of producing FADH2. The H2O2 is broken down to water by catalase. The fatty acyl-CoA intermediates formed during β-oxidation are the same in peroxisomes and mitochondria. Peroxisomes also contain the necessary enzymes for α-oxidation, which are necessary for the oxidation of some fatty acids with methyl branches. Branched-chain fatty acids also require additional enzymatic modification to enter the alpha-oxidation pathway within peroxisomes. Phytanic acid (3,7,11,14-tetramethylhexadecanoic acid), requires additional peroxisomal enzymes to undergo beta-oxidation. Phytanic acid initially forms phytanyl CoA which is then hydroxylated at the alpha carbon by phytanyl CoA hydroxylase (alpha-hydroxylase), encoded by the PHYH gene. The alpha carbon-hydroxyl bond then undergoes two successive rounds of oxidation to pristanic acid. Pristanic acid undergoes beta-oxidation, which produces acetyl CoA and propionyl CoA in alternative rounds. As with peroxisomal beta-oxidation of VLCFAs, this process generally ends when the carbon chain length reaches 6-8 carbons, at which point the molecule is shuttled to the mitochondria by carnitine for complete oxidation to carbon dioxide and water. Figure $31$ shows the steps in the catabolism of phytanic acid (3,7,11,14-tetramethylhexadecanoic acid). Omega-oxidation The omega-oxidation pathway occurs in the endoplasmic reticulum and is used to metabolize larger fatty acids, which given their hydrophobicity might be damaging to cells in high concentrations. In this pathway the fatty acids are metabolized to dicarboxylic acids which increases their water solubility for excretion in the urine. The first step uses cytochrome P450 enzymes that are also used to modify xenobiotic compounds with dioxygen, making them more soluble as well. The omega oxidation pathway is shown in Figure $32$. Three subfamily members of the cytochrome P450s (CYPs) show a preference for hydroxylation of short-chain fatty acids (C7-C10, CYP4B), medium-chain (C10-C16, CYP4A), and long-chain (C16-C26, CYP4F) fatty acids. which can be saturated, unsaturated and branched, fatty acids. Figure $33$ shows a summary of the alpha, beta, and omega oxidation pathway Diseases of fatty acid metabolism Listed below are a few select diseases that either directly involves defective fatty acid metabolism through intrinsic enzyme deficiencies or indirectly prevent the proper functioning of fatty acid metabolism through extrinsic enzyme deficiencies. Many, but not all, deficiencies of enzymes involved in fatty acid oxidation result in abnormal neurological development and or function early in life; a brief list of signs and symptoms appears under the selected diseases mentioned. MCAD Deficiency Medium-chain acyl dehydrogenase is the most common inherited defect of fatty acid oxidation in humans; as one would expect, medium-chain, 6-8 carbon molecules accumulate in this disease. Clinical manifestations of MCAD deficiency primarily present during fasting conditions and include lethargy, weakness, sweating, and hypoglycemia, most commonly in children under the age of 5. Serum measurements of octanoyl carnitine are usually elevated in these patients and can aid in the diagnosis. These abundant molecules then undergo oxidation by the cytochrome P450 system involved in omega-oxidation, resulting in a dicarboxylic acidemia and dicarboxylic aciduria. Zellweger Syndrome Zellweger syndrome results from autosomal recessive mutations in the PEX genes, which code for peroxin proteins needed for the assembly of peroxisomes. Almost 70% of all peroxisomal biogenesis disorders (PBDs) result from a PEX1 gene mutation. Many different fatty acid compounds can accumulate without the oxidative machinery of peroxisomes, including VLCFAs and phytanic acid. Manifestations of this disease generally include the brain, kidneys, and skeleton. X-Linked Adrenoleukodystrophy (X-ALD) X-ALD is a genetic deficiency of the ABCD transporters in the membrane of peroxisomes, as mentioned previously, resulting in the pathological accumulation of phytanic acid and VLCFAs within cells and is most clinically significant when the ABCD1 transporter is absent. The disease presents with neurodegenerative and adrenal abnormalities. Refsum Disease Refsum disease results from a genetic deficiency of the enzyme phytanyl CoA 2-hydroxylase, which, as previously mentioned, is involved in the alpha-oxidation of phytanic acid, a breakdown product of chlorophyll. Notable clinical manifestations of Refsum disease include cardiac malfunction and defective functioning of the olfactory and auditory nerves due to the accumulation of phytanic acid.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/17%3A_Fatty_Acid_Catabolism/17.02%3A_Oxidation_of_Fatty_Acids.txt

Search Fundamentals of Biochemistry Much of the material below derives from Kolb et al. BMC Medicine (2021) 19:313 https://doi.org/10.1186/s12916-021-02185-0. Creative Commons Attribution 4.0 International License, http://creativecommons.org/licenses/by/4.0/. Introduction Ketone bodies are the name given to two molecules, acetoacetate (a ketone) and D-β-hydroxybutyrate (its reduction product) that derive from the condensation of acetate in the form of acetyl-CoA. Their structures are shown in Figure \(1\). They are synthesized when acetyl-CoA concentrations are high. That would occur under a couple of conditions: • fatty acids are being mobilized in adipocytes and broken down into acetyl-CoA through β-oxidation in the mitochondria; • the acetyl-CoA produced can not enter the Kreb cycle leading to the buildup of acetyl-CoA in the matrix. The latter condition arises when carbohydrate metabolism is compromised so the citric acid cycle has slowed. This occurs if the citric acid cycle intermediate oxaloacetate is pulled away from the cycle for the gluconeogenic synthesis of glucose. Pyruvate, another source of acetyl-CoA (through pyruvate dehydrogenase) is also depleted in gluconeogenesis. Of course, we are describing conditions in the liver, the major site of gluconeogenesis, and also of ketone body synthesis. These conditions are also met in the underfed or starving state when glycogen supplies are exhausted and fatty acid reserves are being used for energy. Likewise, these conditions are met in diabetes when glucose might be abundant in the circulation but its entry into the cell is compromised by problems with insulin signaling. Fatty acids released by adipocytes into the blood are bound to serum albumin for transport to tissue. Albumin, with bound fatty acids, does not, however, cross the blood-brain barrier. Ketone bodies are very small and water soluble so they can cross the blood-brain barrier, where they can deliver a soluble equivalent of fatty acids to the brain and other tissues in times of energy need. Ketone bodies can be considered solubilized and easily transportable fatty acid equivalents. They have low pKa values of 3.6 (acetoacetate) and 4.7 (D-β-hydroxybutyrate) so they can lead to "ketoacidosis" in diabetics. Acetone is a nonacidic and sweet-smelling molecule, which can be detected in the breath of diabetics, is produced spontaneously and through catalysis through the decarboxylation of the beta-keto acid acetoacetate. It is also considered a ketone body. Blood-Brain barrier The blood–brain barrier (BBB) protects the entire nervous system (CNS). Endothelial cells line the surface of all blood vessels. In the CNS there are additional structures that prevent the entry of many molecules. Tight and adherence junctions form between endothelial cells that prevent the movement of solutes between the cells. Additional protection is provided by the basement membrane, glial cells, and pericytes. In addition, drug efflux transporters (ABC transporters) move xenobiotics (foreign toxic molecules back through the barrier. Nonpolar molecules like dioxygen molecules and low molecular weight can cross the BBB by simple diffusion. Other molecules like glucose and amino acids are carried across by transporters. Larger substances can move in through receptor-mediated endocytosis. Albumin, the major carrier of free fatty acids, like other proteins, does not readily pass through the barrier, and its drug-macromolecule complex, cannot cross. Figure \(2\) shows a representation of the blood-brain barrier. Figure \(2\): Structure of the blood-brain barrier. Wilhelm et al. Int. J. Mol. Sci. 2013, 14, 1383-1411; doi:10.3390/ijms14011383. Creative Commons Attribution License (http://creativecommons.org/licenses/by/3.0/). In this section, we will explore the synthesis and utilization of ketone bodies. The synthesis pathway is called ketogenesis, and the degradation pathway is called ketolysis. Ketone body production and use are intimately tried to fatty acid metabolism so it is appropriate to discuss them right after fatty acid oxidation. An overview of both ketogenesis and ketolysis is shown in Figure \(3\). Figure \(3\): Overview of the ketogenesis (left) and ketolysis (right) pathways. FFA, free fatty acids; mThiolase, mitochondrial thiolase; HMGCS2, hydroxy methylglutaryl-CoA synthase; HMGCL, HMG-CoA lyase; BDH1, mitochondrial βOHB dehydrogenase; MCT1/2, monocarboxylate transporter 1 and 2; SCOT, succinyl-CoA:3-oxoacid-CoA transferase; CS, citrate synthase. Kolb et al. BMC Medicine (2021) 19:313 https://doi.org/10.1186/s12916-021-02185-0 Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/. Most acetoacetate is converted to its reduction product, β-hydroxybutyrate, for blood transport. It can spontaneously or through enzymatic catalysis, can be converted to acetone since it is a β-ketoacid with a built-in electron sink that stabilizes the negative charge in the transition state and intermediate. When ketone bodies enter cells, the β-hydroxybutyrate is converted back to acetoacetate and eventually to acetyl-CoA. Ketogenesis: The details Figure \(4\) shows the chemical structures of all intermediates in the synthesis of ketone bodies. We'll study three of the reactions in ketogenesis in more detail below. Reaction 1: Thiolase We have presented the mechanism of thiolase in Step 4 of mitochondrial β-oxidation of fatty acid using the enzyme acetyl-CoA acetyltransferase (ACAT1) also called 3-ketoacyl-CoA thiolase. The final step in the beta-oxidation pathway involves cleavage of the bond between the alpha and beta carbon by CoASH. This step is catalyzed by beta-keto thiolase and is a thiolytic (as opposed to cleavage by water - a hydrolysis) reaction. The reaction produces one molecule of acetyl CoA and a fatty acyl CoA that is two carbons shorter. The process repeats until the even chain fatty acid is completely converted into acetyl CoA. The activity of the enzyme is reversible and it can also catalyze the Claisen condensation of two acetyl-CoA molecules into acetoacetyl-CoA, which occurs in the first step of ketogenesis. This step makes a C-C bond. Reaction 2: HMG-CoA Synthase This enzyme catalyzes the condensation of acetoacetyl–CoA (AcAc–CoA) and acetyl–CoA (Ac-CoA) to form 3-hydroxy-3-methylglutaryl (HMG)–CoA, through the formation of a C-C bond. Forming C-C bonds is critical to all biosynthesis. It's not a simple process. Cofactors are often used such as thiamine pyrophosphate, as we saw with pyruvate dehydrogenase. The reverse process, breaking a C-C bond, is also difficult unless the atoms are activated. One way to activate a carbon atom is to oxidize it through hydroxylation, often using dioxygen in reactions catalyzed by cytochrome P450. Likewise, C-C bond formation often occurs through the reaction of radicals in hemolytic reactions or through the rearrangement of key ionic intermediates. Radical reactions can be between a carbon free radical and heme derivatives. Radical reactions involve heme cofactors (as in P450), non-heme iron proteins (where the iron ion can cycle between different oxidation states), flavoproteins (in which the flavin can undergo both 1 and 2 electron redox steps), radical S-adenosylmethionine (SAM) enzymes, and cobalamins. HMG-CoA synthase is somewhat unique in that it forms a C-C bond by activating the methyl group of acetyl-cysteine. The acetyl group comes from an acyl-CoA "donor". The enzyme catalyzes the first committed step in the formation of complex isoprenoids (like cholesterol) and ketone bodies. The product, 3-hydroxy-3-methylglutaryl HMG–CoA, can either be reduced by HMG-CoA reductase to form mevalonate, which leads to cholesterol synthesis, or cleaved by the enzyme HMG-CoA lyase, to produce acetoacetate, a ketone body. Since it catalyzes the first step in cholesterol synthesis, the enzyme HMG-CoA reductase has been a primary focus of drug therapy to reduce high levels of serum cholesterol, which is generally associated with cardiovascular disease. Over 200 million people worldwide are on statins, the primary drugs used to block HMG-CoA reductase activity. Luckily, it doesn't affect HMG-CoA lyase. Gram-positive bacteria also require the mevalonate pathway. HMG-CoA synthase catalyzes a bisubstrate reaction that displays ping-pong kinetics, characteristic of a covalent enzyme intermediate. The first substrate binds to the enzyme and transfers an acetyl group to a nucleophilic Cys 111 in the active site to form an acetyl-Cys 111 intermediae. Free CoA departs. Next the second substrate, acetoacetyl-CoA binds, and condenses with the acetyl group donated by acetyl-Cys 111. This condensation involves an enolate. A plausible reaction mechanism is shown in Figure \(5\). Figure \(5\): Reaction mechanism for HMG-CoA synthase Hence there are three parts of the reaction: acetylation/deacetylation, condensation/cleavage with an enolate intermediate, and C-C formation and hydrolysis/dehydration. Figure \(6\) shows an interactive iCn3D model of the Staphylococcus aureus HMG-COA Synthase with bound HMG-CoA and acetoacetyl-CoA (1XPK) Figure \(6\): Staphylococcus aureus HMG-COA Synthase with bound HMG-CoA and acetoacetyl-CoA (1XPK). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...deiFs7JceE5H76 The biologically active unit (homodimer) is shown with each monomer shown in a different color. The A chain (light cyan) has bound HMG-CoA (HMG) while the B chain (light gold) has acetoacetyl-CoA (CAA) bound. The Glu 79, Cys 111, and His 233 in each subunit are shown in CPK sticks and labeled. Note that the Cys 111 is covalently modified in each subunit. Figure \(7\) shows an interactive iCn3D model of the human 3-hydroxy-3-methylglutaryl CoA synthase I with bound CoASH (2P8U) Figure \(7\): Human 3-hydroxy-3-methylglutaryl CoA synthase I (monomer) with bound CoASH (2P8U). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...xxEoUfecTk3eL6 The active site side chains are numbered differently (Glu 95, Cys 129, and His 264) compared to the S. aureus protein. Only the monomer is shown in this model. Reaction 3: HMG-CoA Lyase The lyase reaction is a C-C bond cleavage to produce acetyl-CoA and acetoacetate. A gene knockout of this gene in mice is lethal. Figure \(8\) shows a plausible mechanism for the reaction. Figure \(9\) shows an interactive iCn3D model of the human HMG-CoA lyase with the competitive inhibitor hydroxyglutaryl-CoA (HG-CoA)(3MP3) Figure \(9\): Human HMG-CoA lyase with inhibitor HG-CoA (3MP3). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...VJBWxXpcKQyij8 Only one subunit of the homodimer is shown. The key residues involved in catalysis and binding (Arg 41, Asp 42, His 233, His 235, and Cys 266) are shown in sticks, CPK colors, and labeled. The competitive inhibitor, 3-hydroxyglutaryl-CoA (without the methyl group) is shown in CPK-colored sticks. The S-stereoisomer of HMG-CoA or inhibitor binds, not the R-isomer. Water may shuttle protons between Asp 42 and the C3 hydroxyl of HMG-CoA. Arg 41 facilitates the proper enol/keto tautomer of the product mutation on reaction product enolization. Mutation of Arg 41 have lhas effect on catalysis. We won't go into detail on the remaining enzymes since we have encountered variants of NAD+/NADH dehydrogenase reactions before. The acetoacetate decarboxylase just hastens the already spontaneous decarboxylation of the β-keto ester acetoacetate. Regulation of Ketogenesis Ketogenesis can be upregulated by hormones such as glucagon, cortisol, thyroid hormones, and catecholamines which lead to fatty acid mobilization. Insulin is the primary hormonal regulator of this process. Insulin regulates many key enzymes in the ketogenic pathway, and a state of low insulin triggers the process. A low insulin state leads to: • Increased free fatty acids (FFAs) arising from decreased inhibition of hormone-sensitive lipase • Increased uptake of FFAs into the mitochondria arising from decreased activation of acetyl-CoA carboxylase, decreasing malonyl CoA, which disinhibits Carnitine Palmitoyltransferase 1 (CPT1) • Increased production of ketone bodies arising from increased HMG-CoA activity Ketolysis: Utilization of ketone bodies Figure \(10\) shows the chemical structures in the utilization of ketone bodies. Since we have encountered reactions similar to these before, we will not discuss their detailed mechanisms. Most organs and tissues can use ketone bodies for energy. They are a major source of energy for the brain when glucose is not available. That makes sense given that fatty acids can't cross the blood-brain barrier. The heart prefers fatty acids for energy but can also use ketone bodies. The liver makes but does not use them since the gene for step 2, beta ketoacyl-CoA transferase is not transcribed. D-β-hydroxybutyrate is first converted to the other ketone body, acetoacetate, for the pathway to continue. The net result is the formation of two acetyl-CoAs that can then enter the citric acid cycle for ATP production. Acetone is a dead-end ketone body and is excreted in the urine or eliminated in the breath. Clinical Significance An overproduction of ketone bodies through increased ketogenesis can cause ketoacidosis (given the pKas of the ketone bodies). One type is diabetic ketoacidosis (DKA) in Type I and II diabetes which impair glucose use. This leads to liver gluconeogenesis and the resulting depletion of oxaloacetate and pyruvate, impacting the ability of acetyl-CoA to enter the citric acid cycle. On presentation, patients are usually very dehydrated from being hyperglycemic. High glucose levels in the blood cause higher osmotic pressure in the vessels and decreased water reabsorption in the kidneys. They often have accompanying symptoms like confusion, nausea, vomiting, and abdominal pain. Because of the acidosis, patients often breathe very deeply and rapidly to eliminate carbon dioxide, which can cause and cause respiratory alkalosis. Ketoacidosis also can occur with severe alcoholism and prolonged starvation. Ketogenic Diet The ketogenic diet, characterized by high fat and low carbohydrate consumption, can lead to weight loss. It shifts metabolism to more closely resemble the fasting state. Its long-term effects are not clear but some have suggested the use of ketone-based drugs to mimic the effect of ketone bodies. Fasting can also lead to a ketogenic state and it has been used for the treatment of epilepsy before the drug became available. It is important to differentiate this attempt to induce a ketogenic state from diabetic ketoacidosis.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/17%3A_Fatty_Acid_Catabolism/17.03%3A_Ketone_Bodies.txt

Search Fundamentals of Biochemistry Introduction Organic chemistry is usually described as the chemistry of carbon-containing molecules. But isn't that definition a bit carbon-centric, especially since the prevalence of oxygen-containing molecules is staggering? What about nitrogen? We live in a dinitrogen-rich atmosphere (80%), and all classes of biomolecules (lipids, carbohydrates, nucleic acids, and proteins) contain nitrogen. Dinitrogen is very stable, given its triple bond and its nonpolar nature. We rely on a few organisms to fix N2 from the atmosphere to form ammonium (NH4+), which through nitrification and denitrification can form nitrite (NO2-), nitrate (NO2-), nitric oxide (NO), and nitrous oxide (N2O), the latter being a potent greenhouse gas. We'll concentrate on the metabolic fate of amino groups in amino acids and proteins in the next section. Before exploring their fates, look at the figure below which shows an overall view of the biological nitrogen cycle. The study of biochemistry should encompass more than homo sapiens and expand to the ecosystem in which we are such a small but damaging part. Let's break down the diagram from a biochemical perspective. There are aerobic and anaerobic processes (conducted by bacteria). Nitrogen-containing substances include both inorganic (ammonium, nitrate, nitrite) and organic (amino acids, nucleotides, etc) molecules. The reactions shown are oxidative and reductive (note: the oxidation number of the nitrogen atoms in the molecules is shown in red). Most of the reactions are carried out underground by bacterial and Archaeal microorganisms. Here are some of the major reactions: • N2 fixation (a reduction): N2 from the air is converted by bacteria to ammonium (NH4+) by the enzyme nitrogenase of soil prokaryotes. The energetically disfavored reaction requires lots of ATPs. Ammonium once made can then be taken up by primary producers like plants and incorporated into biomolecules such as amino acids, which animals consume. For those who may still believe that people have marginal effects on our biosphere, consider this. We may soon fix more N2 (to NH3) through the industrial Born-Haber reaction (used for fertilizer and explosive productions) that all the N2 fixed by the biosphere. Much of the nitrogen in use comes from the Born-Haber reaction. The excess NH4+ (upwards of 50%) produced industrially and which enters the soil in fertilizers (mostly as NH4NO3) has overwhelmed nature's ability to balance the nitrogen cycle and is not taken up by plants. It is metabolized by microorganisms to nitrite and nitrate. • Nitrification: Ammonium is converted to nitrite by ammonia-oxidizing aerobic microorganisms and further to nitrate by a separate group of nitrite-oxidizing aerobic bacteria. Here are the reactions (Rx 1 and 2) to produce nitrate through a hydroxylamine intermediate, followed by the formation of nitrate (Rx 3). $\ce{NH3 + O2 + 2e^{-} -> NH2OH + H2O} \tag{Rx 1}$ $\ce{NH2OH + H2O -> NO2^{-} + 5H^{+} + 4e^{-}} \tag{Rx 2}$ $\ce{NO2^{-} + 1/2 O2 -> NO3^{-}} \tag{Rx 3}$ These added ions exceed soil capacity and end up runoff water, polluting our rivers and lakes. • Denitrification: This anaerobic reaction pathway reproduces N2 from nitrate Here is the net reaction: $\ce{2NO3^{-} + 10e^{-} + 12H^{+} -> 2N2 + 6H2O} \nonumber$ • Anammox reaction: This more recently discovered bacterial anaerobic reaction pathway converts ammonium and nitrate to N2. Here is the net reaction $\ce{NO2^{-} + NH3^{+} -> N2 + 2H2O} \nonumber$ • Ammonification (not to be confused with mummification) occurs when plants and animals decompose, which returns ammonium to the soil for reuse by plants and microbes. These reactions are shown in the abbreviated Nitrogen Cycle below. Nitrogen metabolites are nutrients for plants and perhaps the most important nutrients in the regulation of plant growth (primary productivity) and in regulating life diversity in the biosphere. All living organisms require feedstocks to produce energy and as substrates for biosynthetic reactions. Which ones are used depends on the organism. Plants are primary producers so they use their own synthesized carbohydrates for both energy production and biosynthesis. For carnivores, proteins and their derived amino acids are the source of energy (through oxidation) and serve as biosynthetic precursors. For omnivorous organisms, the source of energy depends on the "fed" state. With abundant food resources, carbohydrates, and lipids are the source of energy. Unlike carbohydrates and lipids, which can be stored as glycogen and triacylglycerols for future use, excess protein, and their associated amino acids can not be stored, so amino acids can be eliminated or used for oxidative energy. In the fed state, carbohydrates are the main source, while in the unfed state, lipids take a predominant role. Under starving conditions, the organisms' own proteins are broken down and used for oxidative energy production and for any biosynthesis that remains. In diseased states like diabetes, which can be likened to a starving state in the presence of abundant carbohydrates, both lipids and amino acids become the sources of energy. How are amino acids in animals oxidatively metabolized? Many pathways could be used to do so but it would seem logical that NH4+ would be removed and the carbons in the remaining molecule would eventually enter glycolysis or the TCA cycle in the form of ketoacids. NH4+ is toxic in high concentrations. Ammonium is not oxidized to nitrite or nitrates in humans as occurs in the soil by microorganisms. It can be recycled back into nucleotides or amino acids, and excess amounts are eliminated from the organism. Both processes must be highly controlled. We will turn out attention to the oxidation of amino acids in the next section.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/18%3A_Nitrogen_-_Amino_Acid_Catabolism/18.01%3A_The_Biochemistry_of_Nitrogen_in_the_Biosphere.txt

Search Fundamentals of Biochemistry From the previous section on the nitrogen cycle, it should be clear that NH3/NH4+ has a central role in metabolism. Nature's fertilizer for plants is ammonium derived from bacterial nitrogenase and human-derived Born-Haber process. Animals get their ammonia mostly from ingested plants (primary producers) and animal protein. In vertebrates, proteins get digested into small proteins and peptides starting in the stomach where pepsin cleaves proteins after aromatic and Leu side chains into smaller fragments. The low pH in the stomach (1-2) facilitates protein unfolding, which allows great access to buried side chains for pepsin cleavage. As the ingested materials move into the small intestine, the protein fragments are further cleaved into small peptides by trypsin, and chymotrypsin and then into individual amino acids by carboxypeptidases (which cleaves C-terminal amino acids) and aminopeptidases (which cleave at the N-terminal end). The individual amino acids are then adsorbed, along with di- and tripeptides into the epithelial cells lining the small intestine. Peptidases cleave short peptides and amino acids are moved into the blood through cell membrane transporters which are coupled to sodium ion intake. Amino acids are taken up by tissues for use in protein synthesis and by the liver of vertebrates for metabolic processing. The gut proteolytic enzymes are secreted into the gut lumen as precursors (proenzymes), where they are activated either autocatalytically or by other proteases into their mature form. Amino acids are also derived from the degradation of cellular proteins by a supramolecular assembly called the proteasome. Proteins designated for cleavage are covalently modified by ubiquitin, a short ubiquitous protein, which targets them to the proteasome. Degradation of amino acids to produce ammonium Here is some good advice on giving a seminar: tell what you will tell them, tell them and tell them what you told them. These words imply that repetition is one of the keys to learning. So look at Figure \(1\) which shows an overall view of the glycolytic and TCA pathways and see how amino acid catabolism fits into what you already know! Figure \(1\): Amino acids and their involvement with the glycolytic and TCA pathways A summarizing principle is that amino acids degrade to form intermediates that fit into these two common pathways and can be used for ATP production as well as biosynthesis. Those that form pyruvate are glucogenic amino acids, while those that form acetyl-CoA are ketogenic (form ketones and could form ketone bodies as well). Many form direct intermediates in the TCA cycle. This chapter will focus on glutamine/glutamic acid, alanine (which forms pyruvate), and aspartic acid. In this section, we will focus on the role of amino acid catabolism in producing ammonium. When new students confront the structures, properties, and reactions of amino acids, they are undoubtedly daunted. Even biochemists would be if amino acid metabolism is outside their active research area. Perhaps some interesting biological properties of free amino acids might pique your interest. For example, did you know that ... • glutamate and glutamine are the most abundant amino acids in red blood cells; • alanine and glutamine are the most abundant amino acid in the body; • glutamic acid, glutamine, and alanine are the most abundant amino acids in the cerebrospinal fluid CSF (50-55% of total amino acids); • glutamine is the most abundant amino acid in blood serum; • In contrast, leucine is the most abundant amino acid in proteins, and free leucine is a prime regulator of protein synthesis through its interaction with the mTOR kinase complex. The other top three amino acids in proteins are serine, lysine, and glutamic acid. From the above list, it appears that Glu and Gln play special roles in metabolism and signaling. They also play major roles in ammonium metabolism as the pair are major sources of NH4+ production in cells. Gln has two Ns (an amine and an amide), so it's not an unexpected source of nitrogen: • Glutamine can give up its amide nitrogen to form NH4+ on conversion to glutamic acid, a reaction catalyzed by glutaminase • Glutamic acid can undergo oxidative deamination to form α-ketoglutarate (a TCA intermediate) and free NH4+, a reaction catalyzed by glutamate dehydrogenase • Glutamic acid can also give up its ammonia nitrogen to a ketoacid like pyruvate to form α-ketoglutarate (a TCA intermediate) and another amino acid (alanine if pyruvate was used as the keto acid), a reaction catalyzed by a class of enzymes called transaminases (makes sense again), which are also called aminotransferases. In this case, free NH4+ is not formed but rather is passed to a keto acid to form another amino acid. In vertebrates, free amino acids are metabolized in the liver. Amino acids that enter the liver transfer their ammonia group to α-ketoglutarate (aka 2-oxoglutarate) to form glutamic acid which enters the mitochondria and can be cleaved by glutamate dehydrogenase to form α-ketoglutarate and free ammonium. The NH4+ produced is either recycled or excreted in the form of NH4+ in fish, urea, H2N(C=O)NH2 in vertebrates, and uric acid in birds and reptiles. Excess amino acids (which again can't be stored for energy) in other organs pass their ammonia group to glutamic acid to form glutamine, which then heads to the liver for processing. Glutamine then becomes a safe way to transfer 1-2 ammonium equivalents through the blood. Alanine and aspartic acid also play secondary roles. Ala is one transamination step away, through the removal of its amine group, from pyruvate, a crucial α-ketoacid end product of glycolysis and entry product for the TCA cycle (after pyruvate dehydrogenase). In muscle, excess amino acids pass their ammonia groups to pyruvate to form alanine, another "safe" carrier of ammonium, which heads to the liver for processing. Likewise, aspartic acid is one transamination step way, through the removal of its amino group, to form oxaloacetate, another TCA α-ketoacid. Figure \(2\) shows the key α-keto acids which are also intermediates of the TCA cycle or feed into it (pyruvate) and their respective transamination amino acid products. Their structures immediately show a link between amino acid and carbohydrate metabolism. Figure \(2\): α-ketoacids and their transamination amino acid products We will focus most of our attention on glutamine and glutamic acid. Figure \(3\) summarizes the important enzymatic steps in the conversion of Gln ↔ Glu ↔ α-KG Figure \(3\): Important enzymatic steps in the conversion of Glutamine ↔ Glutamic acid ↔ α-ketoglutarate Pyridoxal phosphate and transamination reactions Most free amino acids start their metabolic degradation in the liver with transamination reactions, using PLP as a cofactor. PLP enables other biochemical reactions of amino acids, including racemizations, decarboxylation, and dehydration (of serine). PLP is covalently attached to a Lys through a Schiff base linkage in the enzyme during the reaction cycle so it is not considered a substrate, but rather a cofactor that returns to its original state after the reaction. Figure \(4\) shows the structure of PLP and the imine formed on reaction with an amino acid. The reaction is readily reversible through hydrolysis in the presence of water which would presumably not be present in the active site of PLP-dependent enzymes. A lysine ammonia group in the enzyme forms a covalent adduct with PLP to form an imine. The imine in the figure below is called an internal aldimine. Internal implies that the source of the N in the imine link is a Lys internal to the protein. If in your mind you would replace the N in the imine with an O, the functional group with the C=O would be an aldehyde. Hence the imine shown is an aldimine. Figure \(4\): PLP and its imine-protein conjugate William Jencks, in his classic text, Catalysis in Chemistry, describes the mechanistic beauty of PLP-dependent enzymes: "It has been said that God created an organism especially adapted to help the biologist find an answer to every question about the physiology of living systems; if this is so, it must be concluded that pyridoxal phosphate was created to provide satisfaction and enlightenment to those enzymologists and chemists who enjoy pushing electrons, for no other coenzyme is involved in such a wide variety of reactions, in both enzyme and model systems, which can be reasonably interpreted in terms of the chemical properties of the coenzyme. Most of these reactions are made possible by a common structural feature. That is, electron withdrawal toward the cationic nitrogen atom of the imine and into the electron sink of the pyridoxal ring from the alpha carbon atom of the attached amino acid activates all three of the substituents of this carbon for reactions which require electron withdrawal from this atom." When PLP-dependent enzymes react with an external amino acid acting as a substrate, the amine of the incoming amino acid replaces the amine from the enzyme's active Lys to produce an external imine. The figure below shows an external amino acid in imine linkage to PLP, and in the form shown is an aldimine. Each PLP-dependent reaction involves a protonated and positively charged pyridinium N as an electron sink, which facilitates cleavage of each bond around the Cα of the covalently attached amino acid. Bond cleavage would leave a lone pair and negative charge on the α-carbon and no place to stabilize it by resonance. On the formation of a Schiff base with a pKa of around 7.0, the imine nitrogen, especially in its protonated form, is an excellent stabilizer of the negative charge. Continued delocalization of the lone pair and negative charge to the positive pyridinium nitrogen further facilitates the stabilization and hence the reaction. Figure \(5\) shows the cleavage of bonds around the Cα of an amino acid-PLP Schiff base. Figure \(5\): Cleavage of bonds around the Cα of an amino acid-PLP Schiff base Figure \(6\) shows the mechanism for a transamination reaction using PLP as a cofactor. This is of course the reaction most relevant to this chapter section. The amino acid substrate is first shown in an aldimine linkage to PLP. Also, note the conversion of an aldimine to a ketimine in the next step. Figure \(6\): Mechanism for a transamination reaction using PLP as a cofactor The model below shows the active site of aspartate transaminase from E. Coli (pdb code 1aam) with PLP in Schiff base linkage with lysine 258. Three amino acids critical for enzyme function (Trp 142, Asp 223, and Lys 258) and all side chains within 5 angstroms from the active site defined by those amino acids, are shown. Lys 258 forms the Schiff base with PLP cofactor and also acts as a general acid/base on interconversion between the aldimine and ketamine forms (see figure above). The ring of Trp 142 forms pi-stacking interactions with the aromatic ring of PLP and helps position it. Asp 223 acts as a general acid/base in the catalytic cycle, facilitating the deprotonation of the amine group of the substrate Asp, which makes it a potent nucleophile in the formation of the external aldimine. Figure \(7\) shows an interactive iCn3D model of the active site of aspartate transaminase from E. Coli with PLP (1aam) Figure \(7\): Active site of aspartate transaminase from E. Coli with PLP (1aam). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...YeQ6kah4EuERT9 The various names and abbreviations for transaminases/aminotransferases (which are reversible) can be confusing. Two key ones whose levels are used to assess liver toxicity in clinical tests are AST (GOT) and ALT (GPT). • ASpartate Amino Transferase (AST) is the same enzyme as Glutamate Oxalacetate Transaminase (GOT), named for the reverse reaction: aspartate + α−ketoglutarate ↔ oxalacetate + glutamate • ALanine Amino Transferase (ALT) is the same enzyme as Glutamate Pyruvate Transaminase (GPT), named for the reverse reaction: alanine +α−ketoglutarate ↔ pyruvate + glutamate We will consider other reactions of PLP-dependent enzymes when needed. To summarize, different amino acids coming into the liver can donate their amine groups to α-ketoglutarate to form glutamic acid in a reaction catalyzed by a transaminase/aminotransferase to form glutamic acid. Production of NH3 - Glutamate Dehydrogenase Now that we made glutamic acid from various amino acids, we can now break it down to make α-ketoglutarate again and ammonium. The overall reversible reaction catalyzed by the enzyme glutamate dehydrogenase in mitochondria is shown in Figure \(8\). Mammalian livers can also use NADP+ as an oxidizing agent. Figure \(8\): Overall reversible reaction catalyzed by the enzyme glutamate dehydrogenase in mitochondria Humans have two isozymes. GluDH1 is expressed at high levels in the liver, brain, pancreas, and kidney, while GluDH2 is found in the retina, testes, and brain. GluDH1 can use both NAD+ and NADP+ so it can be used in both anabolic and catabolic reactions. The reaction/product pairs again show this reaction to be a clear link between protein and carbohydrate metabolism. Figure \(9\) shows just the first step in the reaction of glutamate dehydrogenase. Figure \(9\): First step in the reaction of glutamate dehydrogenase The C-N in Glu becomes C=N in the imine intermediate. You should recognize this as an oxidation step even if the obvious oxidizing agent, NAD(P)+, was not shown. That should also be evident since a hydride (H-) with a lone pair is removed from glutamic acid and not a H+ as in an acid/base reaction. Subsequent steps include nucleophilic attack by water (enhanced by a proximal Lys acting as a general base) on the imine C followed by ammonia release on the formation of alpha-ketoglutarate. The overall reaction is an oxidative deamination. NH3/NH4+ are toxic in high concentrations. One possible reason is that the NH4+ cation might compete with the transport of other ions across neural membranes, altering transmembrane potentials and hence neural function. Given this toxicity, you expect that glutamate dehydrogenase is highly regulated and it is in mammals. The mammalian enzyme is a hexamer of identical subunits which suggests allosteric regulation of activity (much like tetrameric hemoglobin). ADP and leucine are allosteric activators while GTP, palmitoyl CoA, and ATP act as inhibitors. The GTP/ADP regulation is consistent with the notion that the enzyme is regulated by the need for cellular energy (remember that GTP is formed in the TCA cycle). Although this reaction is written above in the direction of NH3 formation, the reaction is reversible, even though the ΔG0=-6.2 kcal/mol (-26 kJ/mol) and though the human enzymes have a high Km for ammonia (12-62 mM, Brenda Database). As the pH decreases from 8 to 7, the Km for ammonia increases from around 12 to 60 mM. This implies that on glutamate uptake and in acidotic conditions, the reaction runs exclusively in the direction of NH3 release - as an oxidative deamination reaction. Transporting ammonium equivalents in the blood - Glutamine and Alanine NH3/NH4+ are toxic in high concentration, mechanisms must be in place to transport it in the blood. Ideally, it would be transported in a nontoxic form. Glutamic acid is a key metabolic molecule so glutamine is the preferred molecule for safe "NH3/NH4+" transport from tissue to the liver, while alanine is the choice for muscle tissue. In a way, this less toxic mode of transport is similar in principle to the use of soluble ketone bodies to transport the energy equivalent of the less soluble fatty acids in the blood. A key enzyme for this process is glutamine synthetase, which catalyzes this reaction: NH4+ + ATP + Glu → Gln + ADP + HPO42- + H+ This reaction is important as most ingested glutamine is converted to glutamate on absorption. NH3/NH4+ made in the intestine and kidney ends up in the circulation where it heads to the liver for the eventual production of urea. Figure \(10\) shows the Glucose-Alanine Cycle in comparison to the Cori cycle we discussed in a previous chapter. Figure \(10\): Glucose-Alanine Cycle in comparison to the Cori cycle Production of NH3 - Glutaminase Most amino acids arriving in the liver for degradation go through two enzymes, transaminases to form glutamic acid followed by glutamate dehydrogenase to form a TCA intermediate (alpha-ketoglutarate) and ammonia. Glutamine is a carrier of excess ammonia in the bloodstream (see below). When it arrives in the liver, it can lose its amide NH2 as ammonium as it is converted to glutamic acid by the enzyme glutaminase, a nonoxidative enzyme that is expressed in the liver, brain, and kidney. Some of the glutamate can lose ammonia by the enzyme glutamate dehydrogenase, but most is reserved for protein synthesis or the creation of anabolic precursors. As mentioned above, glutamine is the most abundant free amino acid in the body and most cells. It can donate its amide N through reactions catalyzed by amidotransferase (not to be confused with aminotransferases) leading to the incorporation of nitrogen into many biomolecules. Metabolic Summary • Glutamine is both a carrier of ammonia and a carbon backbone used in metabolism when converted to α-ketoglutarate. It can be metabolized for energy and used in biosynthesis for nucleotides and neurotransmitters. As such its levels are controlled by glutamine synthase and glutaminase. When we get to synthetic pathways, you will find that glutamine is a NH3/NH4+ donor for the synthesis of other amino acids as well as nucleotides, amino sugars, and NAD+. • Glutamine that is absorbed in the intestines is mostly converted to glutamic acid in the intestinal epithelial cells and is transported by the blood. The rest heads to the liver for processing. Hence most glutamine must be synthesized by glutamine synthase, a cytosolic enzyme found in most mammalian cells. However, it is most abundant in muscle, liver, and adipose cells which express little glutaminase and from which it can be exported. Glutaminase in the liver and kidney leads to ammonium production • Glutamine is used as a source of energy and carbon for biosynthesis by tumor cells, which need both energy and intermediates for synthesis by rapidly proliferating cells. Aerobic glycolysis ("Warburg effect") allows that to occur.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/18%3A_Nitrogen_-_Amino_Acid_Catabolism/18.02%3A_Metabolic_Fates_of_Amino_Groups.txt

Search Fundamentals of Biochemistry The Urea Cycle Now we are in a position to see what happens to excess NH3/NH4+ that accumulates in the liver mitochondria. The ammonia that is formed in the liver through oxidative deamination by glutamate dehydrogenase, ends up in urea. Alternatively, it could be added to glutamic acid to form glutamine which can head to the kidney where, after sequential reactions by glutaminase (a deamidation reaction) and glutamate dehydrogenase, it forms ammonia for direct excretion in the urine. If your diet is high in proteins and hence amino acids, since they can't be stored as proteins per se, they are metabolized for energy and the metabolic products can be converted to fat or carbohydrate. That leaves excess nitrogen which ends up mostly as urea for excretion. Urea is a very water soluble, nontoxic molecule that biochemists use in the lab to chemically denature protein (at concentrations of 3-6 M). Clinical blood urea concentrations are expressed as Blood Urea Nitrogen (BUN) which range from 7 to 20 mg/dL N, which is equivalent to 2.5 to 7.1 mM urea. Serum levels depend predominately on the balance between urea synthesis in the liver and its elimination by the kidney. Normal cellular urea concentrations should be similar. Urea is produced by a pathway called the urea cycle, as shown in Figure \(1\). Figure \(1\): The Urea Cycle The enzyme names are: • CPS1: carbamoyl phosphate synthase I • OTC: ornithine transcarbamoylase • ASS1: arginosuccinate synthase 1 • ASL: arginosuccinase lysase (aka arginosuccinase) • ARG1: arginase It is color-coded to designate the sources of the C and N atoms. One N comes from ammonia (or indirectly from the amido N of glutamine), while the C atom comes from carbonate. The other comes from the amine of aspartate. Note that 3 ATPs are used to power the cycle. Also note the guanidino group of arginine (NH(NH2+)NH2) is the immediate donor of the 2 N atoms in the step that produces urea. Knowing that one amino acid was the immediate donor of the N atoms in urea, you would easily predict that. Now, let's look at some of the steps, starting with the ones that are powered by ATP, which are also the ones where nitrogen is incorporated into the precursors of urea. Carbamoyl phosphate synthase I (CPS I) This enzyme catalyzes the first "committed" path of the pathway and makes the reactant that enters the cycle, carbamoyl phosphate. It requires a specific cofactor, N-acetyl-glutamate (NAG), produced by the acetylation of glutamate by acetyl-CoA using an enzyme NAG synthase which is activated by arginine. The reaction is not part of the cycle but provides the input for it. A cytosolic version of this enzyme, CPS II, is used to synthesize arginine and pyrimidine nucleotides using glutamine as a donor of NH4+which reacts with carboxyphosphate to produce carbamoyl phosphate. It is not regulated by NAG. CPS I provides a way to form a (C=O)NH2 unit for urea by condensing bicarbonate and ammonia to form a carbamate, which contains a high energy "motif" with respect to its hydrolysis product. (Note: this does not imply that the broken bond is high energy, a misnomer found in many books.) Hence the reaction is powered by 2 ATPs as an energy source. Urea is a molecule that "carries" both NH3/NH4+ and also carbonate. A reaction mechanism for carbamoyl phosphate synthase is shown in Figure \(2\). Figure \(2\): Reaction mechanism for carbamoyl phosphate synthase The two high-energy motif molecules (again with respect to their hydrolysis products) are protected from nonspecific hydrolysis as the mixed anhydride pass along a sequestered tunnel to a more distal phosphorylation site in the multimeric enzyme. Figure \(3\) shows an interactive iCn3D model of the human carbamoyl phosphate synthetase I with bound ADP and N-acetyl-glutamate (5DOU). Figure \(3\): Human carbamoyl phosphate synthetase I with bound ADP and N-acetyl-glutamate (5DOU). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...yV8fW3mW9idy6A The model shows one chain of the multimer. The 2 ADPs and single N-acetylglutamate (NLG) are shown in spacefill and labeled. Additional bound ions are shown (unlabeled). The protein exists in a monomer-dimer equilibrium mixture. Each monomer has 1 NAG and 2 ADP, which represent the two different binding sites for the two different phosphorylation steps as shown in the reaction above. The two phosphorylation sites are connected by a tunnel allowing passage of the mixed anhydride to the carbamate phosphorylation site. The NAG is an allosteric modifier as it is not bound near the phosphorylation sites. On binding to the apo form of the enzyme, it elicits a large conformation change allowing a competent enzyme conformation with a complete tunnel to predominate. An image of the tunnel for one monomer of the human carbamoyl phosphate synthetase I is shown in Figure \(4\). Figure \(4\): Tunnel for one monomer of the human carbamoyl phosphate synthetase I enzyme. Two ADPs are shown below in spacefill with CPK color. The tunnel between the two ADP sites is highlighted in magenta. The yellow molecule is NAG, which again exerts its activation effect by binding to an allosteric site, which effectively opens up the tunnel. Figure \(5\) shows the enzyme monomer in a different orientation with a tunnel made using ChExVis: a tool for molecular channel extraction and visualization. The large red and green spheres show the surface openings. Figure \(5\): Second view of the tunnel in human carbamoyl phosphate synthetase I The top panel in Figure \(6\) shows the radius in Angstroms of the tunnel going from the Red sphere to the Green sphere in the image above. The bottom panel shows a measure of the hydrophobicity (red)/hydrophilicity (blue) of the amino acids surrounding the tunnel. Figure \(6\): Tunnel size and polarity profile in human carbamoyl phosphate synthetase I ASS1: ArginoSuccinate Synthase 1 Another ATP is cleaved to drive the conversion of citrulline to arginosuccinate, as shown in Figure \(7\). The product could easily be cleaved in the next step to form the amino acid arginine. Figure \(7\): ATP cleavage drives the conversion of citrulline to arginosuccinate ASL: arginosuccinase lysase (aka arginosuccinase) This enzyme catalyzes a beta-elimination reaction, which may proceed through either an E2 or E1 mechanism. A general but very abbreviated mechanism showing a two-step (E1) elimination proceeding through a carbanion intermediate is shown in Figure \(8\): below. Figure \(8\): Mechanism for conversionof arginosuccinate to arginine and fumarate Figure \(9\) shows an interactive iCn3D model of the tetrameric arginosuccinate lyase (ASL) from Mycobacterium tuberculosis (6IEN). Figure \(9\): Tetrameric arginosuccinate lyase (ASL) from Mycobacterium tuberculosis (6IEN). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...32EuouGQ6xMVk6 Each monomer of the four monomers is shown in a different color. 3 arginosuccinates (substrate) and 1 fumarate and 1 arginine (products) are shown in spacefill, with CPK colors and labeled. Each monomer has an N-, M-, and a C-domain. The four binding sites consist of residues from three monomers. The products are in the fourth site. Catalytic residues probably include a serine and histidine act as general acids/bases with a deprotonated lysine making the serine a general base. Ornithine Transcarbamyolase The reaction mechanism here is quite simple. The carbamoyl phosphate is an activated electrophile in which the carbonyl C is attacked by a deprotonated amino of ornithine to produce citrulline. The gene is found on the X chromosome in humans so mutations in males lead to a "hyperammonemic coma" and death. Heterozygous females can be asymptomatic and that generally proves to be fatal. Heterozygous females are either asymptomatic or have problems arising from defects in pyrimidine biosynthesis which a low-protein diet can alleviate with the addition of arginine. Arginase There are two general variants of this enzyme, Arginase I is a cytosolic enzyme expressed in the liver and is involved in urea synthesis. Arginase II is a mitochondrial enzyme and is involved more generally in arginine metabolism. Arginase I converts arginine to ornithine and urea.  The enzyme has 2 Mn ions in the active site.  A hydroxide ion bridges the two Mn ions and acts as a nucleophile which attacks the guanidino group of arginine and forms a tetrahedral intermediate. An active site Asp 128 acts as a general acid and protonates the amino leaving group to form ornithine and urea.  Water then adds back to the Mn dinuclear cluster and ionizes, donating a proton to solvent through an intermediary His 41 as it reforms the active enzyme. The model below shows the trimeric human arginase I (3KV2) in complex with the inhibitor N(omega)-hydroxy-nor-L-arginine, in which one of the guanidino Ns is replaced with an OH. The two grey spheres are Mn ions. Figure \(10\) shows an interactive iCn3D model of the trimeric human arginase I in complex with the inhibitor N(omega)-hydroxy-nor-L-arginine (3KV2). Figure \(10\): Trimeric human arginase I in complex with the inhibitor N(omega)-hydroxy-nor-L-arginine (3KV2). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...QaAQpt5t2KKic6 Figure \(11\) show an active site water (#480) near the Mn ions. Figure \(11\): Active site of human arginase I with an active site water Water molecules can act as ligands and bind transition state metal ions through a coordinate covalent bond. The pKa of the bound water shifts to a lower value, making it more likely to deprotonate to form OH-, which is both a better base and nucleophile. Water 480 in the active site appears to form a coordinate covalent bond to the Mn ion forming an active site hydroxide that attacks the guanidino group of arginine, leading to the formation of urea and ornithine. ​Regulation If excess amino acids/proteins are consumed, a change in gene expression can increase levels of the enzymes in the urea cycle. In the short term, the activity is regulated by the enzyme that provides access into the cycle, carbamoyl phosphate synthase, whose activity is regulated by N-acetylglutamic acid and which catalyzes the rate-limiting step. As shown above, without the allosteric regulator, NAG, the enzyme is functionally inactive. The levels of NAG are determined by the enzyme NAG synthase, which itself is regulated by free arginine, the immediate precursor of urea in the cycle. Supplemental arginine is given to patients with urea disorder system and its effect probably occurs through the regulation of NAG synthase. The link between the urea cycle and the TCA cycle Two of the major pathways you have explored, the TCA and urea cycles, are not linear but cyclic pathways. What advantages do "circular" offer over linear ones? Perhaps a comparison to economic pathways offers a clue. In a linear economy, raw materials are used to produce a product which is eventually discarded as trash. Unfortunately, linear pathways dominate our world at a great cost to our environment. To reduce the inefficiencies in a linear economy and make it more viable and to decrease environmental damage and associated climate change, the world should move to a circular economy. A circular economy is characterized by what has been referred to as the 3Rs: reduce, reuse, and recycle. This saves resources and energy. These characteristics also apply to circular biochemical pathways as reduced levels of metabolites get reused and recycled. Given this, it should not come as a surprise that the urea and TCA cycle are linked, especially given that fumaric acid is a product of the urea cycle and a cyclic metabolite of the TCA cycle. In addition, aspartic acid is one transamination step away from oxaloacetate. The interconnections between the urea cycle and the TCA cycle are shown in Figure \(12\) along with two additions, the malate aspartate shuttle, and the aspartate arginosuccinate shunt. Figure \(12\): Urea and TCA cycles linked by the malate aspartate shuttle and the aspartate arginosuccinate shunt. One of the nitrogens in urea comes from the amine of aspartate, which can be formed through the transamination of oxaloacetate from the TCA cycle to form aspartate. Since the TCA cycle is a cycle, an equivalent of one oxaloacetate must return to the cycle. It does so indirectly from fumarate, a TCA intermediate produced as a by-product of the urea cycle, but the fumarate is produced in the cytoplasm. There, it gets converted to malate, which can get transported into the mitochondria by a transport protein, mitochondrial 2-oxoglutarate/malate transporter, that is part of yet another cycle shown in the figure above, the aspartate arginosuccinate shunt. The introduction of fumarate into the shuttle occurs through the aspartate arginosuccinate shunt shown in the figure above. Once malate enters the mitochondrial matrix, it can be directly converted to oxaloacetate. Note that these linked cycles require the presence of both mitochondrial and cytoplasmic forms of the several enzymes and balances between two key amino acids across the mitochondrial divide, glutamate, and aspartate. One such key enzyme is ASpartate Amino Transferase (AST), aka Glutamate Oxaloacetate Transaminase (GOT), named for the reverse reaction of aspartate + α−ketoglutarate ↔ oxaloacetate + glutamate. As mentioned in the previous chapter section, this key enzyme can be found in the blood if the liver is damaged, so its levels are routinely used in medical tests for impaired liver function. Another key function of the malate aspartate shuttle is to bring into the mitochondria reduced "equivalents" of NADH that are produced in the cytoplasm from glycolysis and fatty acid oxidation. There is no membrane transporter for NAD+ or NADH. Instead, cytoplasmic malate, produced by the cytosolic reduction of oxaloacetate by NADH, can be transported into the matrix which can reform mitochondrial NADH on reconversion to oxaloacetate. The NADH can then be used to power ATP production through mitochondrial electron transport/oxidative phosphorylation pathways. From an energetic perspective, four phosphoanhydrides bonds in 3 ATP molecules are used to produce one urea molecule. This large energy expenditure is partially compensated for by ATP made from the "fumarate equivalents" that enter the mitochondria through the aspartate arginosuccinate and aspartate arginosuccinate shunts and the passage of NADH through oxidative phosphorylation.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/18%3A_Nitrogen_-_Amino_Acid_Catabolism/18.03%3A_Nitrogen_Excretion_and_the_Urea_Cycle.txt

Search Fundamentals of Biochemistry Amino acid degradation We saw how nitrogen is removed from amino acids to produce urea or NH4+ in the previous chapter section. What are the fates of the carbon skeletons that remain? This section is where students might get overwhelmed by the diversity of amino acid degradation pathways, so it helps to realize that carbon skeletons of deaminated amino acids can be used for biosynthesis or energy production and are converted to key glycolytic and TCA intermediates that you have seen many times before. Everything is interconnected which makes the study of metabolism daunting but also fascinating, as organisms try to extract all the energy and molecule-building atoms from a metabolite and minimize waste. Here are some key features of amino acid catabolism that this chapter section will present. • some are converted to pyruvate, the end product of glycolysis and the start reactant of gluconeogenesis. Hence, these amino acids are glucogenic; • some are converted to acetoacetate-CoA and or acetyl-CoA. Both of these can be converted to ketone bodies (acetoacetate/β-hydroxybutyrate) so these are considered ketogenic. Since the two carbons of the acetyl group of acetyl-CoA are lost as CO2 in the TCA cycle, and there is no reverse for the pyruvate dehydrogenase reaction (pyr → acetyl-CoA), acetyl-CoA formed by amino acid degradation can not be used to create glucose in net fashion; • some are metabolized to form TCA intermediates. Since they are added in a net fashion to the TCA cycle and don't remove the existing pool of TCA intermediates, they can produce in a net fashion directly or indirectly molecules that can be used to produce glucose. These entry reactions to the TCA which replenish or add to TCA intermediates are called anaplerotic (replenishing) reactions. Hence these amino acids are also glucogenic. • some have multiple ways to be degraded and can produce both acetyl-CoA and pyruvate, so they are both glucogenic and ketogenic. Let's get more explicit: • purely ketogenic: only Leu and Lys (the only amino acids whose name starts with L and you have to Love them since there are only 2 amino acids in this category) • both: 5 including the aromatics - Trp, Tyr, Phe - and Ile/Thr • purely glucogenic: the rest General hints as to if an amino acid is glucogenic and/or ketogenic can be derived from their structures. Fully or Partly Ketogenic Amino Acids: Amino acids except for Arg that have a continuous chain of 3 or more carbon atoms in their side chains and hence are more "fat-like" are ketogenic or ketogenic/glucogenic. These include Lys, Leu, Ile, and the aromatic amino acids Phe, Trp, and Tyr and exclude Val. (note: Thr, which is both ketogenic and glucogenic, doesn't fit this rule). The fully or partly ketogenic amino acids are shown in Figure \(1\). Purely Glucogenic Amino Acids (for Pyr and/or TCA intermediates: Which amino acids produces which intermediate? Except for glycine, it's pretty easy to remember. The number of carbons in the intermediate formed is the same as the number of carbons in the longest chain in the amino acid. These amino acids form the 5C TCA intermediate alpha-ketoglutarate (2-oxoglutarate) and are shown in Figure \(2\). These amino acids form either the 4C TCA intermediate succinate or oxaloacetate. Amino acids with more oxidized 4C atoms in the continuous chain produced oxaloacetate (which is more oxidized than succinate), while the least oxidized ones produce succinate acid (which is more reduced than oxaloacetate). These amino acids are shown in Figure \(3\). The 3C amino acids, Ser, Ala, and Cys, except for glycine, form pyruvate, as shown in Figure \(4\). A full figure summarizing everything above is shown in Figure \(5\)! With a little help from my friends: Cofactors and Amino Acid Catabolism The myriad of breaking, making, and rearranging of carbon atoms in amino acid catabolism is daunting. As with all reactions, a pathway for a flow of electrons and stabilization of transition states and intermediates must be in place for the reactions to occur. All methods of catalysis (general acid/base, covalent/nucleophilic catalysis, electrostatic/metal ion catalysis, preferential stabilization of the transition state occur. Yet some reactions need additional "helpers" or "cofactors" to facilitate electron flow and shuttle small motifs (from electrons in redox reactions to methyl groups in methylases) from one molecule to another. We've seen the important role of pyridoxal phosphate in transaminations/aminotransferases in the previous section. Even the terminology cofactor is a bit confusing since analogous terms are used in different contexts: • cofactors - nonprotein small molecules or ions (divalent ions, transition metal ions, and Fe/S clusters for example) that must bind to an enzyme, but once bound usually stay put. That is, they don't dissociate during the catalytic cycle. For instance, PLP must initially bind to an enzyme but then becomes covalently attached through a Schiff base through the epsilon amino group of a Lys in the catalytic site. That linkage might swap with an incoming amino acid substrate, for example, but reforms in the catalytic cycle so the enzyme remains functional. • coenzyme - a vitamin derivative organic cofactor, as compared to inorganic ions, for example. Coenzyme is a poor historical name that persists. • prosthetic group - this is usually a coenzyme that is either covalently bound (like PLP) or noncovalently bound with such a low KD (like FAD/FADH2-containing enzymes) that they stay bound • cosubstrate - these bind as substrate (for example NAD+), and depart as products (for example NADH). Here is a table of common nonmetallic cofactors. Cofactor Vitamin Derivative Carrier biotin H - biotin 1 C - CO2 (most oxidized) tetrahydrofolic acid (FH4 or THF) B9 - folic acid 1C - formyl, -(C=O)H, methylene (-CH2) (more reduced) and methyl -CH3 cobalamin B12 - cobalamin methyl CH3 thiamine pyrophosphate B1 - Thiamine 2C group coenzyme A B5 - pantothenic acids acetyl (CH3-C=O) and acyl (R-C=O) pyridoxal phosphate B6 - pyridoxine amino and carboxyl groups NAD+/NADP+ B3 - Niacin electrons FAD/FMN B2 - riboflavin electrons Examples of other nonvitamin cofactors include S-adenosylmethionine (SAM or adoMet), a carrier of methyl groups, coenzyme Q, a carrier of electrons, tetrahydrobiopterin, a carrier of oxygen and electrons, and of course heme, a carrier of electrons. We've already described amino acid deamination reactions using PLP. Now let's consider the biochemistry of two of these cofactors involved in 1 C transfers in amino acid metabolism in more detail, tetrahydrofolate (FH4) and SAM. Tetrahydrofolate (FH4) Tetrahydrofolate (FH4) is a key cofactor in the metabolism of amino acids but it is also critical in the biosynthesis of nucleotides. It is also commonly abbreviated as THF, especially in the names of enzymes that use it. To avoid confusion with chemist's use of THF for tetrahydrofuran, it will be referred to mostly here as FH4 but the enzymes will be named as derivatives of the abbreviation XHF (X = D for dihydro and T for tetrahydro. FH4 is a carrier of 1 C units in various oxidation states. It receives 1C units and then transfers them to other species. The structure of FH4 and derivatives carrying 1 C units with different oxidation numbers (indicating their oxidation state) are shown in Figure \(6\). FH4 is made from the vitamin folate (bacteria, fungi, and most plants can synthesize it), which gets converted first to dihydrofolate (FH2), and then tetrahydrofolate (FH4), using NADPH as a reducing substrate "cofactor", by the enzyme dihydrofolate reductase. The model below shows this small enzyme with NADPH (NADP+) and folate bound to DHFR (7dfr). Folate (FOL) is shown in salmon space fill while NADPH is shown in cyan. Figure \(7\) shows an interactive iCn3D model of dihydrofolate reductase with bound NADPH (NADP+) and folate (7dfr). Figure \(7\): Dihydrofolate reductase with bound NADP+ and folate (7dfr). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...ojdpBZuQH8Lm96 Folate (FOL) and NADP+ (NAP) are shown in spacefill and CPK colors. The side chain of Asp 27 is shown in sticks and CPK colors. The oxygen atoms of two water molecules interacting with folate are shown as red spheres. A disordered and hence mobile loop (residues 16-20) that is involved in binding the nicotinamide is shown in cyan. As FH4 is a key substrate in many reactions, including nucleotide synthesis (as we shall see later), this enzyme is a key target for chemotherapy to kill cancer cells which require robust nucleotide synthesis for rapid cell proliferation. One such drug, which inhibits the enzyme, methotrexate, is shown in Figure \(8\). Before getting into the nitty-gritty of amino acid degradation and tetrahydrofolate chemistry, let's take a look at the mechanism by which dihydrofolate reductase, DHFR, catalyzes the sequential reduction by 2 NADPH molecules of folate to tetrahydrofolate. This is illustrated in Figure \(9\). One Carbon Chemistry: The Interconversion of FH4 1C derivatives Tetrahydrofolate is a carrier for 1C units in metabolism. As such, is it intimately involved in many anabolic and catabolic reactions. These include thymidine and purine biosynthesis and amino acids metabolism through reactions involving serine, glycine (both with 1C in their side chains), and methionine (with 1C after the sulfur in the side chain). The reactions take place in both the cytoplasm and mitochondria. FH4 is involved directly or indirectly in epigenetic control of DNA expression as well, as methylation of both DNA and histones is critical to gene expression. The myriad of 1C derivatives of FH4 make the biochemistry complex, but as the transfer of 1C is a critical job in both the breakdown of the carbon skeleton of amino acids and biosynthesis, we need to explore it. The complexity is simplified by noting that the 1C derivatives have only three different oxidation states (+2, 0, and -2), as noted in Figure \(10\). 1C units usually enter FH4 as the N5,N10 methylene unit. This is made in both the cytoplasm and mitochondria since 1C-derivatized FH4 appears not to cross the mitochondrial membranes. The methylene derivative, with a 1C oxidation # of 0, can be reduced to form methyl (oxidation # -2) or oxidized to methenyl or formyl groups (oxidation # +2). We will see some of these reactions again in chapters dealing with amino acid and nucleotide biosynthesis. That's not a bad thing - learning occurs best on the repetition of material in different contexts. Serine Hydroxymethyltransferase (SHMT): A complex reaction needed two cofactors - PLP and FH4 Let's look in detail at one mechanism that shows how a 1C methylene is added to tetrahydrofolate (FH4). The mechanism shown is for serine dehydratase, aka serine hydroxymethyltransferase. The enzyme not only uses tetrahydrofolate as a substrate but also PLP, which, as we have seen previously, makes bonds to the alpha-carbon of amino acids labile to cleavage. In this case, the amino acid serine becomes dehydrated through an alpha-elimination reaction. Here is the overall reaction. Serine + FH4 ↔ Glycine + N5,N10-CH2-FH4 + H2O Figure \(11\) shows the dehydration reaction and formation of glycine. using PLP as a cofactor. Figure B shows how the released formaldehyde reacts with FH4 to form N5,N10-methylene FH4, using FH4 as a cofactor. As this is needed in purine and thymidylate synthesis, SHMT is a target for malaria treatment as well. Glu 57 plays a key role as a general acid/base throughout the catalytic cycle of the enzyme. Pathway diagrams showing a myriad of reactants, products, and enzymes can be confusing to students (and to authors as well). It's useful to think about them and see them in different ways. Here is another way to present the conversion of FH4 and its various 1C intermediates. Figure \(12\) concentrates on the outputs of 1C FH4 derivatives (shown in blue eclipses). The enzymes catalyzing these reactions are shown in the table below. This figure applies to both catabolic and anabolic reactions using FH4 derivatives. Oxidation numbers for the 1C adducts are shown. Any reaction that involves a change in redox state must use NAD(P)+/NAD(P)H as a redox reagent. • N5,N10-methylene FH4 gives thymidine and serine • N5-methyl FH4 gives methionine • N10-formyl FH4 gives formate, purines, and CO2 • N5-formyl serves more as a passive reservoir of 1C units. Table \(1\): Enzymes catalyzing interconversions of THF derivatives Abbreviation Enzyme Name ALDH 10-formyltetrahydrofolate((aldehyde) dehydrogenase. DHFR Dihydrofolate reductase MTHFD methylenetetrahydrofolate dehydrogenase MTHFD1 C-1-tetrahydrofolate synthase, cytoplasmic MTHFD1L monofunctional tetrahydrofolate synthase, mitochondrial MTHFD2/L methylenetetrahydrofolate dehydrogenase 2/2-like MTFMT mitochondrial methionyl-tRNA formyltransferase MTHFR methylenetetrahydrofolate reductase MTR methionine synthase TYMS thymidylate synthetase A key enzyme in these reactions, methylene-THF reductase (MTHFR), irreversibly removes 1C from the cycle depicted above as it forms N5-methyl FH4. As this reaction is a reduction, it requires a reducing agent (NADPH in yeast and animals and NADH in plants). This irreversible removal would deplete the cycle shown so it is allosterically regulated (inhibited) by another methylating agent, S-adenosylmethionine (SAM aka adoMet), as we will see next. Plant versions of the enzyme are reversible so there is no need for regulation by SAM in this feedback loop process. S-adenosylmethionine (SAM), also known as adoMet N5-methyl FH4 appears to have one function, to methylate a molecule called homocysteine (same as Cys but with an extra -CH2 in the side chain) to methionine. This is adenosylated (not phosphorylated!) with ATP to produce S-adenosylmethionine (SAM), a more potent methylating agent than N5-methyl FH4. SAM is hence part of a cycle involving N5-methyl-FH4. The methyl group of N5-methyl FH4 reacts with homocysteine to produce methionine, catalyzed by the enzyme methionine synthase, which requires cobalamin (vitamin B12) as a cofactor. The combined Figure \(13\) adds the Met and Folate cycles (showing the outputs of the 1C Folate metabolism cycle Figure \(14\) shows the structures of molecules in the Met Cycle. The mechanism of methyl transfers using SAM as the -CH3 donor involves a SN2 attack by a nucleophile of the substrate on the CH3 of SAM, with the electron pair from the C-S bond going to the positively charged sulfonium sulfur, a great "electron sink". An analogous nucleophilic attack on the terminal CH3 of plain old methionine would not be readily enabled. Hence SAM, with its charged S, has a much high methyl transfer potential than N5-CH3-FH4. In the actual reaction catalyzed by methionine synthase (MS) in mammals, the methyl CH3 from N5-methyl FH4 is first transferred to cobalamin, a derivative of vitamin B12, to form methylcobalamin, which then transfers it to homocysteine. The structure of the C-terminal half of B12-dependent Methionine Synthase from E. Coli with bound adenosylhomocysteine bound (3iva) is shown below and in this link: Figure \(15\) shows an interactive iCn3D model of the C-terminal half of B12-dependent Methionine Synthase from E. Coli with bound adenosylhomocysteine bound (3iva) Figure \(15\): C-terminal half of B12-dependent Methionine Synthase from E. Coli with bound adenosylhomocysteine bound (3iva). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...E7jTsdvVvx1M1A In mammals, vitamin B12 is a key cofactor in only two enzymes, one being methionine synthase. If vitamin B12 is lacking, N5-methyl FH4 builds up, and since the enzyme that converts N5,N10-methylene FH4 to N5-methyl FH4, methylenetetrahydrofolate reductase (MTHFR), is irreversible. This leads to megaloblastic anemia as precursors to red blood cells that can't mature. Folate deficiencies also lead to anemia. The Serine Glycine One Carbon (SGOC) Metabolic Cycle Figure 12 shows the coupled Folate and Methionine cycles that emphasize the intermediates involved in the 1C -CH3 transfer reaction, an important part of amino acid metabolism and other anabolic and catabolic reactions as well. Interpreting metabolic figures is complicated. Each is designed to emphasize certain selected features. Another way to present Figure 12 is to emphasize metabolites involved in 1C chemistry in general. Figure \(145\) shows what has been called the Serine Glycine One Carbon (SGOC) metabolic cycle. It is just a redrawn version of Figure 12 with attention drawn to non-FH4 molecules involved in 1C transfers, namely serine, glycine, and also formate. This cycle and its key substrates, serine, and glycine, integrate many metabolic pathways and controls the conversion of serine and glycine into outputs essential for other pathways. We will see this cycle again in the chapter on the biosynthesis of amino acids. The pathway is especially important in tumor cells, which need precursors for nucleic acid, protein, and lipid synthesis.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/18%3A_Nitrogen_-_Amino_Acid_Catabolism/18.04%3A_An_overview_of_amino_acid_metabolism_and_the_role_of_Cofactors.txt

Search Fundamentals of Biochemistry Introduction In previous sections, we saw how nitrogen is removed from amino acids to produce urea or NH4+, that some amino acids are glucogenic, ketogenic, or both, and the role of tetrahydrofolate derivatives and S-adenosylmethionine in 1C transfer reactions. Now we can focus on how the carbon skeletons of amino acids are processed during degradation. Here are some key features of amino acid catabolism that were discussed in the previous section. • some are converted to pyruvate, the end product of glycolysis and the start reactant of gluconeogenesis. Hence, these amino acids are glucogenic; • some are converted to acetoacetate-CoA and or acetyl-CoA. Both of these can be converted to ketone bodies (acetoacetate/β-hydroxybutyrate) so these are considered ketogenic. Since the two carbons of the acetyl group of acetyl-CoA are lost as CO2 in the TCA cycle, and there is no reverse for the pyruvate dehydrogenase reaction (pyr → acetyl-CoA), acetyl-CoA formed by amino acid degradation can not be used to create glucose in net fashion; • some are metabolized to form TCA intermediates. Since they are added in a net fashion to the TCA cycle and don't remove the existing pool of TCA intermediates, they can produce in a net fashion directly or indirectly molecules that can be used to produce glucose. These entry reactions to the TCA which replenish or add to TCA intermediates are called anaplerotic (replenishing) reactions. Hence these amino acids are also glucogenic. • some have multiple ways to be degraded and can produce both acetyl-CoA and pyruvate, so they are both glucogenic and ketogenic. Let's get more explicit: • purely ketogenic: only Leu and Lys (the only amino acids whose name starts with L and you have to Love them since there are only 2 amino acids in this category) • both: 5 are, including the aromatics - Trp, Tyr, Phe - along with Ile/Thr • purely glucogenic: the rest Figure \(1\), also shown in the previous sections, summarizes the fates of the 20 amino acids in their catabolic reactions Figure \(1\): Fates of the 20 amino acids in their catabolic reactions Given the myriad of enzymes and pathways involved, we won't delve into the mechanisms for the reactions or the structures of the enzymes, with the exception of one for lysine metabolism. Conversion to Pyruvate: Ala, Trp, Cys, Ser, Gly, Thr We ended section 18:3 with a discussion of the Ser Gly One Carbon Cycle (SGOC), so some of this will be a bit of a review. Figure \(2\) shows an overview of the conversion of amino acids to pyruvate. More details are provided for each of the steps below. Figure \(2\): Overview of conversion of amino acids to pyruvate The metabolic steps for the chemical transformations shown in A-F are described in more detail below. Tryptophan to alanine and on to acetoacetate This is a multistep process as shown in Figure \(3\). Figure \(3\): Conversion of Tryptophan to Alanine and to acetoacetate The starting material, tryptophan, is highlighted in a red box while the end product of specific interest, Ala, is highlighted in a green box. No reaction occurs in isolation in a cell, but rather as part of a more complex pathway. In the figure below, Ala is presented almost as a side product as the modified aromatic ring found in either anthranilate or 3-hydroxyanthranilate continues on to form either acetoacetate, a ketone body which can breakdown to acetyl-CoA (making tryptophan ketogenic as well as glucogenic) or NAD+. Alanine to Pyruvate As described in 18.2, and shown in Figure \(4\), the PLP-dependent enzyme ALanine Amino Transferase (ALT), also known as Glutamate Pyruvate Transaminase (GPT), catalyzes this simple transamination reaction: alanine +α−ketoglutarate ↔ pyruvate + glutamate Figure \(4\): Alanine to pyruvate The glutamate produced in this reaction can be oxidatively deaminated to give NH4+ and α-ketoglutarate again, giving the net reaction: Threonine to Glycine There are several pathways for this conversion. One involves the conversion of Thr to 2-amino-3-ketobutyrate by threonine-3-dehydrogenase. Rx: Thr + NAD+ ↔ 2-amino-3-ketobutyrate + NADH This is followed by the conversion of 2-amino-3-ketobutyrate to glycine by the enzyme 2-amino-3-ketobutyrate coenzyme A ligase. Rx: 2-amino-3-ketobutyrate + CoASH ↔ Gly + acetyl-CoA The net of these reactions is Rx: Thr + NAD+ + CoASH ↔ Gly + acetyl-CoA + NADH These reactions are illustrated in Figure \(5\). Figure \(5\): Threonine to glycine A second and predominate reaction involves the conversion of Thr to NH4 + and α-ketobutyrate by the PLP-dependent enzyme Ser/Thr dehydratase (also called threonine ammonia-lyase), an enzyme we have seen in the previous section. Note this reaction does NOT produce glycine but is an intermediate, α-ketobutyrate. Rx: Thr ↔ NH4 + + α-ketobutyrate α-ketobutyrate can then be converted to propionyl-CoA. Rx: α-ketobutyrate + NAD+ + CoASH ↔ propionyl-CoA + NADH + CO2 + H+ This reaction, catalyzed by the inner mitochondrial membrane branched-chain α-ketoacid dehydrogenase complex (BCKDC or BCKDH complex) is an oxidative decarboxylation reaction. BCKDC is a member of two other enzymes, pyruvate dehydrogenase and alpha-ketoglutarate dehydrogenase, both of which act on short alpha-keto acids to produce key Kreb cycle metabolites. Propionyl CoA is then converted eventually in several mitochondrial steps to succinyl CoA for entrance into the TCA cycle. Three enzymes are required for this conversion: propionyl CoA carboxylase, methylmalonyl-CoA epimerase, and methylmalonyl-CoA mutase. Propionyl carboxylase, like another alpha-keto acid carboxylase (pyruvate carboxylase), requires ATP, Biotin, and CO2 (as a substrate) for the carboxylation reaction and hence is often referred to as an ABC enzyme. The three-step conversion pathway of propionyl CoA to succinyl CoA is also used for in the degradation of Valine, Odd-chain fatty acids (which form multiple 2-carbon acetyl CoA units and 1 3-C propionyl CoA unit), Methionine, and Isoleucine along with Threonine. This three-step pathway is sometimes referred to as VOMIT pathway. The third pathway, which we just saw in the previous section, is catalyzed by serine hydroxymethyltransferase (SHMT) (but also called glycine hydroxymethyltransferase or threonine aldolase) and requires the use of both PLP and tetrahydrofolate as cofactors. A 1C methylene is added to tetrahydrofolate (FH4). PLP makes bonds to the alpha-carbon of amino acids labile to cleavage. In this case, the amino acid threonine becomes dehydrated through an alpha-elimination reaction. However, threonine has an extra CH3 group which is released as acetaldehyde. Here is the overall reaction. Rx: Thr+ FH4 + ↔ Glycine + N5,N10-FH4 + acetaldehyde + H2O The enzymes involved in this reaction are a bit unclear in the literature. It appears that SHMT can act on Thr at a lower rate, but that a second enzyme, threonine aldolase, which seems to be afunctional in mammals, acts in other organisms. Glycine to Serine As mentioned above, this reversible reaction is catalyzed by serine hydroxymethyltransferase (SHMT) (see the mechanism in section 18.4) and uses tetrahydrofolate and PLP as cofactors. Here is the overall reaction, the reverse of the Gly ↔ Ser we saw in 18.4. Rx: Glycine + N5,N10-CH2-FH4 + H2O ↔ Serine + FH4 Figure \(6\) shows the serine dehydratase reaction presented in Chapter 18.4 Figure A below shows the dehydration reaction and formation of glycine. using PLP as a cofactor. Figure B shows how the released formaldehyde reacts with FH4 to form N5,N10-methylene FH4, using FH4 as a cofactor. Figure \(6\): Reversible reaction of Serine to Glycine Serine to Pyruvate This reaction is analogous to the Ala → Pyr reaction in Rx B above and is catalyzed by the PLP-dependent enzyme serine/threonine dehydratase/threonine deaminase. Rx: Serine ↔ Pyr + NH4+ The enzyme is found in the cytoplasm and is mainly involved in gluconeogenesis. Cysteine to Pyruvate The overall reactions for this conversion are shown in the figure below. The aspartate aminotransferase used in the production of 3 sulfinylpyruvate is cytosolic and not the same as the more abundant version in the mitochondria. The reaction pathway is shown in Figure \(7\). Figure \(7\): Cysteine to pyruvate Other important metabolites are made from cysteine catabolic pathways. One is taurine, which is the most abundant free amino acid in the body and is especially abundant in development and early milk. It is synthesized predominately in the liver. It is unclear if hypotaurine is converted to taurine in a non-enzymatic fashion or by an oxidase/dehydrogenase. The sulfate produced in the pathways is used to make an interesting derivative of ATP, 3′-phosphoadenosine-5′-phosphosulfate (PAPS), which is used to produce sulfated sugars using in glycolipid and proteoglycan synthesis. This is illustrated in Figure \(8\). Figure \(8\): Sulfate conversion to PAPS Conversion to Acetyl-CoA: Trp, Lys, Phe, Tyr, Leu, Ile, Thr An overview of the many reactions in ketogenic amino acid degradation is shown in Figure \(9\). The red-boxed amino acids are those that form either acetoacetate (a ketone body) or acetyl-CoA directly (green boxes). Some of the carbons are color-coded red or green to indicate where they end up. Figure \(9\): Ketogenic amino acid pathway Trp to acetyl-CoA Fortunately, we have explored the conversion of the non-ring part of tryptophan to alanine and a precursor of acetoacetyl Coa (2-amino-3-carboxymuconate 6-semialdehyde - ACMS) and to NAD+ (quinolinate). ACMS, through the action of ACMS decarboxylase leads to acetoacetyl CoA and then to acetyl-CoA as shown in Figure \(10\). As Trp is a ketogenic amino acid, it seems appropriate to show the steps that lead to acetyl-CoA even at the risk of providing too much detail. Figure \(10\): Part 2 - Tryptophan to acetyl-CoA Lys metabolism In the liver, the main pathway (of several) starts with the formation of saccharopine from the transamination reaction of lysine and α-ketoglutarate, allowing the ε-amino group of lysine to enter the nitrogen metabolic pool. This transamination does not use pyridoxal phosphate (PLP). The first two steps of the reaction are catalyzed by an enzyme, α-aminoadipic semialdehyde synthase, with two activities (condensation/reduction and hydrolysis/oxidation). Lysine is an essential amino acid since the transamination is not reversible. Figure \(11\) shows pathways for the conversion of lysine to acetoacetyl-CoA and acetyl-CoA. Figure \(11\): Pathways for conversion of lysine to acetoacetyl-CoA and acetyl-CoA. The lysine-oxoglutarate reductase (LOR) and saccharopine dehydrogenase (SDH) are found in one bifunctional enzyme, often called aminoadipic semialdehyde synthase. Figure \(12\) shows an interactive iCn3D model of the AlphaFold predicted structure of aminoadipic semialdehyde synthase (Q9UDR5) Figure \(12\): AlphaFold predicted structure of aminoadipic semialdehyde synthase (Q9UDR5). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...Pi1c7Y5A9DmLa8 The cyan domain represents the lysine-oxoglutarate reductase (LOR) domain. It is connected by a well-predicted alpha helix to the magenta saccharopine dehydrogenase (SDH) domain. Phenylalanine conversion to Tyrosine and continues to acetoacetate We'll follow the conversion of phenylalanine to tyrosine, which continues on to acetoacetate, making Phe and Tyr both ketogenic amino acids, and in subsequent steps that produce fumarate. They can enter the TCA cycle leading to the net production of oxaloacetate, which can be pulled off into gluconeogenesis, making both Phe and Try glucogenic as well. A new cofactor that facilitates electron flow in the conversion of Phe to Try in the first step, catalyzed by the enzyme tyrosine hydrolase, is required. That cofactor is tetrahydrobiopterin (BH4). The reaction involves the hydroxylation of BH4 and then its transfer to phenylalanine. Figure \(13\) shows a possible mechanism for the conversion of phenylalanine to tyrosine with tetrahydrobiopterin (BH4). Figure \(13\): Conversion of phenylalanine to tyrosine with tetrahydrobiopterin (BH4) As in the case with the conversion of dihydrofolate back to tetrahydrofolate (FH4) by dihydrofolate reductase, the 4a-OH-BH4 is converted to dihydrobiopterin and then to tetrahydrobiopterin by dihydrobiopterin reductase. Figure \(14\) shows the full pathway for the conversion of Phe and Tyr to acetoacetate and fumarate. Figure \(14\): Conversion of phenylalanine and tyrosine to acetoacetate and fumarate Leu to Acetoacetate The conversion of leucine to acetoacetate is shown in Figure \(15\). Figure \(15\): Conversion of leucine to acetoacetate The first reaction is a transamination using the PLP-dependent branched-chain aminotransferase (BCAT) with α-ketoglutarate. Isoleucine to Acetyl-CoA Figure \(16\) shows the pathway for the conversion of isoleucine to acetyl-CoA. Figure \(16\): Pathway for conversion of isoleucine to acetyl-CoA. Conversion to α-ketoglutarate: Pro, Glu, Gln, Arg,His Proline and Arginine The conversion of proline (bottom left) to glutamate (top left) is shown in Figure \(17\). Figure \(17\): Conversion of proline (bottom left) to glutamate (top left) Glutamate can then form α-ketoglutarate so the reaction is glucogenic. The conversions of arginine (and proline) to α-ketoglutarate are shown in Figure \(18\). Figure \(18\): Conversion of arginine and proline to α-ketoglutarate Histidine The conversion of histidine to α-ketoglutarate is shown in Figure \(19\). Figure \(19\): Conversion of histidine to α-ketoglutarate As described in the reactions above, can be converted to α-ketoglutarate through transamination reactions. Also, we described in a previous section how glutamine can be deaminated through the action of glutaminase to form glutamine which can likewise form α-ketoglutarate, a gluconeogenic intermediate. Conversion to succinyl-CoA: Met and the branched-chain amino acids Ile, Thr, Val We just saw that two branched-chain amino acids, Leu and Ile, are converted to acetyl-CoA and hence are ketogenic (E and F above). Another branched chain hydrophobic amino acid, Val, and also Leu again, can be converted to succinyl-CoA which can be converted to α-ketoglutarate in the Kreb's cycle in net fashion and hence are glucogenic amino acids. We saw in the introduction to amino acids that produce acetyl-CoA that threonine and isoleucine, two branched-chains amino acids, also form propionyl-CoA which goes on to succinyl CoA. So, let's consider Val, another branched-chain amino acid before we consider Met, both of which have 3 Cs in their side chains. Valine The conversion of valine to succinyl-CoA is shown in Figure \(20\). Figure \(20\): Conversion of valine to succinyl-CoA The other two amino acids with branched-chain carbon chains (isoleucine and leucine), use the same enzymes as valine to enter the degradation pathway. They start with branched-chain transaminases (BCATc or BCATm) followed by oxidative decarboxylation reactions catalyzed by branched-chain ketoacid dehydrogenase (BCKD). Three different enzymes are required for the following dehydrogenase reaction. These are short/branched-chain acyl-CoA dehydrogenase (SBCAD) for isoleucine, isovaleryl-CoA dehydrogenase (IVD) for leucine, and isobutyryl-CoA dehydrogenase (IBD) for valine. Methionine Methionine can be metabolized to S-adenosylhomocysteine (SAM) and on to cysteine and α-ketobutyrate, which can also be produced by a transsulfuration reaction, the produces cysteine. That product is metabolized using branched chains dehydrogenases to eventually produce succinyl-CoA, a TCA intermediate. Three enzymes are needed to convert the α-ketobutyrate to succinyl-CoA. Propionyl-CoA carboxylase uses ATP, biotin, and CO2 while the methylmalonyl-CoA mutase requires vitamin B12. An addition enzyme is an epimerase reaction in which D-methylmalonyl-CoA into L-methylmalonyl-CoA The conversion of propionyl-CoA to succinyl-CoA also occurs for branched-chain amino acids (Val, Ile, Thr) as well as Met, and in addition Odd number fatty acids. This odd assortment of substrates for conversion to succinyl-CoA leads to the name VOMIT pathways. These reactions are illustrated in Figure \(21\). Figure \(21\): Methionine conversion to succinyl-CoA Finally, Aspartate and Asparagine Asparagine is converted to NH3 and aspartate using the enzyme asparagine. Aspartate is then used in a transamination reaction to form oxaloacetate, a gluconeogenic precursor. Aspartate participates in the urea cycle as a way to eliminate nitrogen. Glutamate also acquires NH3 through the reaction catalyzed by glutamine synthase. Both amino acids are substrates for transamination reactions to produce TCA intermediates. Glutamate dehydrogenase can lead to alpha-ketoglutarate. Glutamate and aspartate are important in collecting and eliminating amino nitrogen via glutamine synthetase and the urea cycle, respectively. The catabolic path of the carbon skeletons involves simple 1-step aminotransferase reactions that directly produce net quantities of a TCA cycle intermediate. The glutamate dehydrogenase reaction operating in the direction of 2-oxoglutarate (α-ketoglutarate) production provides a second avenue leading from glutamate to gluconeogenesis.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/18%3A_Nitrogen_-_Amino_Acid_Catabolism/18.05%3A_Pathways_of_Amino_Acid_Degradation.txt

Search Fundamentals of Biochemistry An Overview of Mitochondrial Electron Transport The main oxidizing agent used during aerobic metabolism in the citric acid cycle is NAD+ (although FAD is used in one step). In the process, these oxidizing agents get reduced to form NADH (and FADH2). Unless NAD+ is regenerated, glycolysis and the citric acid cycle will grind to a halt. This occurs under anaerobic conditions when NADH formed in glycolysis is reoxidized back to NAD+ by pyruvate which is converted to lactate. Under aerobic conditions, we are continually breathing one of the best oxidizing agents around, dioxygen. NADH is oxidized back to NAD+ not directly by dioxygen, but indirectly as electrons flow from NADH through a series of electron carriers to dioxygen, which gets reduced to water. This process is called electron transport. No atoms of oxygen are incorporated into NADH or any intermediary electron carrier. Hence the enzyme involved in the terminal electron transport step, in which electrons pass to dioxygen, is an oxidase. The enzymes of the citric acid cycle and electron transport are localized in mitochondria, which is shown in Figure $1$. By analogy to the coupling mechanism under anaerobic conditions, it would be useful from a biological perspective if this electron transport from NADH to dioxygen, a thermodynamically favorable reaction (ΔG of about -55 kcal/mol, -230 kJ/mol), were coupled to ATP synthesis. It is! For years scientists tried to find a molecule with "high energy" (with respect to its hydrolysis product) phosphorylated intermediate similar to 1,3-bisphosphoglycerate (formed by glyceraldehyde-3-phosphate dehydrogenase in glycolysis), which could drive ATP synthesis in the mitochondria. (Remember, there is no such thing as a "high energy" bond.)  None could be found. A startling hypothesis was put forward by Peter Mitchell, which was proven correct and for which he was awarded the Nobel Prize in Chemistry in 1978. The immediate source of energy to drive ATP synthesis was shown to come not from a phosphorylated intermediate, but a proton gradient across the mitochondrial inner membrane. All the enzyme complexes in electron transport are localized in the inner membrane of the mitochondria, as opposed to the cytoplasmic enzymes of glycolysis. A pH gradient is formed across the inner membrane in respiring mitochondria. In electron transport, electrons are passed from mobile electron carriers through membrane complexes back to another mobile carrier. There are four inner mitochondrial membrane complexes involved in the flow of electrons to dioxygen. The first, Complex I, passes two electrons from NADH, which engages like NAD+ in 2 electron redox steps), to a flavin derivative, FMN, covalently attached to Complex I. The oxidation of NADH occurs, as expected, by hydride transfer. The reduced form of FMN then passes electrons in single electron steps (characteristic of FAD-like molecules, which can undergo 1 or 2 electrons transfers) through the complex to the lipophilic electron carrier, ubiquinone, UQ as shown in Figure $2$. This then passes electrons through Complex III to another mobile electron carrier, a small protein, cytochrome C. This protein has a covalently attached heme with a central iron ion that can undergo one-electron redox reactions (Fe3+ + 1e- ↔ Fe2+). Reduce cytochrome C (Fe2+) passes electrons through Complex IV, cytochrome C oxidase, to dioxygen to form water. At each step, electrons are passed to better and better oxidizing agents, as reflected in their increasing positive standard reduction potential. Hence the oxidation at each complex is thermodynamically favored. 4 electrons must be added to dioxygen, along with 4H+ to form water. $\ce{O_2 + 4e^{-} + 4H^{+} → 2H_2O}. \nonumber$ What happened to Complex II? Complex II (also called succinate:quinone oxidoreductase) contains the citric cycle enzyme succinate dehydrogenase, which catalyzes the oxidation of succinate to fumarate by FAD. The bound FADH2 does not dissociate from the enzyme, which is functionally dead for further formation of fumarate until it is oxidized back to FAD by additional components of Complex II. In this process, electrons from FADH2 are passed to the same lipophilic mobile electron carrier from Complex I, ubiquinone. Reduced ubiquinone then can transfer electrons to cytochrome C through Complex III. For Complex I, III, and IV, the energy released by the oxidative event is used to drive protons from the matrix to the intermembrane space of the mitochondria. The oxidative energy released is NOT used to form a "high-energy" mixed anhydride as we saw in the glyceraldehyde-3-phosphate dehydrogenase reaction from glycolysis. The resulting proton gradient collapses in a thermodynamically favorable process that energetically drives the endergonic synthesis of ATP by the last inner membrane complex, ATP synthase, which is sometimes called Complex V. This aerobic coupling of electron transfer (from NADH to dioxygen) and ATP synthesis, is shown schematically in Figure $3$. The electron transfer is usually called electron transport, which might suggest to you that it requires energy as in the case of active transport. It does not, however, since it is strongly favored thermodynamically with dioxygen used as the final electron acceptor. You may wish to question the need for such a complicated series of reactions to move the simple electron from one chemical species to another and a simple H+ from one side of a membrane to another to form a proton gradient that collapses anyway. The movers of these species are aligned in the membrane to optimize the processes and allow the vectorial, nonrandom flow of electrons and protons. This is what evolution has produced. One reason must be to maintain optimal control over the efficiency of the coupled reaction. Another powerful explanation is to reduce dangerous side reactions. In redox chemistry, this involves the formation of highly reactive free radicals that can react quickly and uncontrollably with any species nearby. This is especially true when dioxygen is involved, as we saw in Chapter 13.4 - The Chemistry and Biology of Dioxygen. It takes 4 electrons to fully reduce dioxygen to water. If this process is interrupted, highly reactive oxygen species (ROS) like superoxide and peroxide can form, which can damage membranes, proteins, and nucleic acids. Before we explore each complex in more detail, let's get a lower resolution view of the entire mitochondrial electron transport and ATP synthesis (often called oxidative phosphorylation or ox-phos) which is shown in cartoon form in Figure $4$. Especially note: • the subunit complexity of each Complex; • the location of the subunits (many integral membrane proteins, others more hydrophilic and located in the polar matrix); • the mobile electron carriers NADH/NAD+, lipophilic UQH2/UQ used for both complex I and II, cytochrome C (a protein), and O2; • the number of H+s moved from the matrix to the intermembrane space through Complexes I, III, and IV and back through ATP synthase. Also note that the prokaryotic structures are displayed, even though they are placed in the mitochondrial membrane! Bacteria do not have organelles like mitochondrial but they have the same complexes inserted into their plasma membranes. For bacteria, protons are ejected into the periplasm space between the plasma membrane and cell wall. Diagrams like the one shown in Figure $4$ are really useful in simplifying complex biological systems. At the same time, they constrain our ideas about how, in this case, the individual complexes are arranged in the membrane. Much evidence shows that Complex I, III, and IV (those that pump protons) form a very large supercomplex called the respirasome, which has some 80 subunits in mammals. Clustering the subunits in a supercomplex increases the likelihood that shared mobile electron carriers will not diffuse away, but rather continue passing electrons along the electron transport chain. As mentioned earlier, some reactions generate reactive oxygen species, which need to be kept in the complex to avoid their toxic effects. The stability of the individual complexes is also most likely increased in the larger aggregate. Using cryoelectron microscopy (cryoEM), the structures of two large super- and mega-complexes have been solved showing the arrangement of all of the respiratory complexes. Two different structures showing the SuperComplex-I1III2IV1 (SCI1III2IV1) and MegaComplex-I2III2IV2 (MCI2III2IV2) are shown below in Figure $5$. Panel (A) shows human SCI1III2IV1 (PDB ID: 5XTH) in side view (along the inner membrane) and a top-down view. Note that CI is adjacent to CIII (located in the center) which is adjacent to CIV. This is the expected orientation given the flow of electrons from NADH to dioxygen. In panel (B) substrates and products are included into the side view, along with the flow of H+ (from all of the complexes) into the matrix. Panel C shows the added cofactors (FeS clusters, FMN, hemes, etc). Panel (D) shows a very beautiful top-down view of the human megacomplex, which has two copies of each of subunit. Again note the centrality of Complex III. Figure $6$ shows another model of the megacomplex with Complex II (CII, which does not translocate protons across the membrane) added. It also shows a tetramer of Complex V (ATP synthase) With this low-resolution background view of the entire mitochondrial oxidative phosphorylation pathway done, now let's explore each pathway in more detail. Complex I - NADH-quinone oxidoreductase Complex I is located in the inner mitochondrial membrane in eukaryotes and in the plasma membrane of bacteria. Bacteria have only 14 subunits while the mammalian Complex I has 45, including the core 14 found in bacteria. Another 31 are term supernumerary (in excess of the "normal number") subunits that support the function of the core. The significance of the extra mammalian subunits is still unclear. A cartoon model with the actual crystal structure of the Thermus thermophilus (bacterial) complex is shown in Figure $7$. The hydrophilic or peripheral domain catalyzes electron transfer while the membrane domain (encoded by mitochondrial DNA) is involved in the active transport of protons. It is one of the biggest enzymes around, so it's exciting to try to understand it. Figure $8$ shows an interactive iCn3D model of the mammalian respiratory complex I (5LDW) Figure $8$: Mammalian respiratory complex I (5LDW). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...xVjRPLLgaLCi17 (both load slowly given the size of the structures). For the external link, use the mouse scroll to zoom the structure to see the FeS clusters and FMN, all of which are labeled. The red and blue parallel "dummy" atoms represent the boundary of the mitochondrial inner membrane. Electron Transfer in Complex I Electron flow occurs from NADH to UQ through a series of one-electron carriers in the hydrophilic or peripheral domain of Complex I. The initial handoff of electrons occurs to a flavin cofactor, FMN, and then through a series of Fe/S clusters, to ubiquinone. These electron acceptors include tetranuclear (Fe4S4) and binuclear (Fe2S2) iron-sulfur clusters, a FMN flavin mononucleotide, and a Mn (II) ion. These are shown in Figure $9$. The tetranuclear Fe4S4 cluster is based on the cubane structure with Fe and S occupying alternating corners of a square in a tetrahedral geometry. Each Fe is also coordinated to thiolate (RS-) from coordinating cysteines and also sulfide (S2)-anions. The actual structure is a distorted cube as shown in Figure $10$, along with that of the binuclear Fe2S2 cluster, whose bond angles also deviate from those in a perfect tetrahedron. Many possible micro-redox states with different standard reduction potentials are possible for tetranuclear Fe4S4 clusters, much as polyprotic acids have multiple pKa values. The two relevant to the tetranuclear clusters in Complex I are shown below: a. Fe(II)Fe(III)3/S2(CysS)21- + e- ↔ Fe(II)2Fe(III)2/S2(CysS)22- (less positive standard reduction potential) b. Fe(II)2Fe(III)2/S2(CysS)22- + e- ↔ Fe(II)3Fe(III)/S2(CysS)23- (more positive standard reduction potential) pKas and E0 values You have studied Bronsted acids in many courses and know the factors that affect the pKa of different acids that are structurally similar. These include the initial charge on a potential proton donor. The relative strength of a series of structurally similar acids can be predicted by examining the factors that stabilize the negative charge on the resulting conjugate base. The more stable the conjugate base, the more acidic the parent acid. Factors that stabilize a negative conjugate base are electronegativity of the atom holding the negative charge, resonance which might delocalize the charge, inductive/electron release effect of substituents, and the hybridization of the atom holding the charge (sp anions are more stable due to more s character with electrons pulled more closely to the nucleus). What about a single acid? The pKa of given acid is a constant for a given set of conditions (temperature, solvent polarity, etc) but it can be altered by changing conditions. The diagram below in Figure $11$ shows ways to "tip" the pKa of a given acid (in this case a simple carboxylic acid) to a lower value, making the acid stronger in the new set of conditions. The same ideas apply to the strength of a potential oxidizing agent (electron acceptor) as described by the E0 value (higher for stronger oxidizing agents/electron acceptors). Figure $12$ shows the factors that could alter the E0 value for a binuclear FeS cluster. In this case, changes in coordinating ligands, analogous to changes in substituents for an acid) are included since most students have probably spent more time thinking about pKa values than E0 values. In either case, it can be understood through Le Chatelier's principle. The equilibrium can be shifted to the right (stronger acids with lower pKa or stronger oxidizing agent/electron acceptor with a higher E0) by preferentially stabilizing products compared to the initial reactants. Electrons are passed singly to oxidized UQ in one electron steps to form UQH2. Fe-S clusters are synthesized predominately in the mitochondria where they serve as redox cofactors in electron transport as described above. They are ubiquitous in all life forms and serve other roles in addition to redox cofactors per se as they serve structural roles in proteins and are used in redox signaling within the cell as they change oxidation states. Many proteins that interact with DNA (repair enzymes, polymerases, and helicases) contain an FeS cluster. Evidence suggests that they placed critical roles in the abiotic evolution of life in the absence of oxygen as a terminal electron acceptor in exergonic oxidation reactions. When oxidation became available, they became potentially became toxic to the cell as the Fe2+ could participate in reactions (such as the Fenton reaction) leading to the generation of deleterious reactive oxygen species (such as superoxide). To prevent toxicity, when delivered to the cytoplasm and nucleus they are carried and delivered by cytoplasmic iron-sulfur assembly (CIA) proteins. Figure $13$ shows models of the complex and paths for electron flow. Electrons "tunnel" from one cofactor to another in Complex I through quantum mechanical "hops". Since all of the redox cofactors after the favored hand-off of electrons from NADH to FMN are FeS clusters, which would have similar E0 values (modulated only by local environments), there is no significant thermodynamic barrier to electron flow. The final N2 FeS cluster has a E0 value about 100 mV higher than the started ones so it most readily accepts an electron. The separation of the hydrophilic mitochondrial matrix domains, where electron transfer occurs, from the transmembrane domains, where proton transfer occurs, is an elegant design that keeps the key and reactive particles, e- and H+, from reacting with each other. The electrons stay on one side of the membrane, as the protons move to the other side (until the return in the last step when Complex V produces ATP). Proton Transfer in Complex I Proton transport occurs in the membrane domain. Available evidence suggests that 4 protons move from the cytoplasm to the periplasmic space against a concentration gradient during a catalytic cycle of Complex I in bacteria and other organisms. Figure $14$ shows multiple views of both the key electron transfer step and H+ ejection from the matrix into the intermembrane space. The key to proton transports is the discrete membrane domains shown in the bottom left part of Panel (a). There are 4 such domains involved from left to right, each one associated with the transport of 1H+. The individual membrane domains have different names and abbreviations depending on the organism. They are from left to right: • Nqo12, NuoL or ND5 • Nqo13, NuoM, or ND4 • Nquo14, NuoN or ND2 • Nqo8, NuoH, or ND1. The Nqo8, NuoH, or ND1 domain is proximal and closely linked to the site for the reduction of UQ to UQH2. The NuoN/Nqo14/ND2, NuoM/Nqo13/ND4 and NuoL/Nqo12/ND5 domains act as and are structurally similar to antiporters. Note also a very long helix that crosses the left three-most antiporter domains. This helix runs parallel to the membrane and probably acts as a lever allowing coupled conformational changes of the three H+ transporters. We looked in a previous chapter at transporters and showed in both cartoon and structural models that they alternated between inward and outward-open conformations. Figure $15$ shows more details of the antiporter domains and the unique structural features allowing proton transport. Each antiporter domain has two 5-helix bundles that display pseudo-symmetry with transmembranes helices number TM4-8 and TM9-13. The next to last helix in each bundle (7 and 9) is interpreted, with unbroken parts of the helices number TM7a/b and TM12a/b. Running along the middle of each of the antiporter-like domains of the membrane domain is a series of conserved charged amino acids (aspartate, glutamate, lysine, and histidine) that participate in a hydrogen bonding network. Molecular dynamics simulations show water molecules (panel b), represented by red dots (for the oxygen atoms), transiently occupying a hydrogen-bonded network of waters that serve as a conduit for H+ flow. An unbound "naked" proton (H+) would not persist long in any aqueous biological system. Panel C shows the conformational flexibility in broken interrupted helices that presumably gates the proton pore open and closed. (a) The membrane domain of complex I have conserved buried charged/hydrophilic residues. NuoL (red); NuoM (blue); NuoN (yellow); NuoH (green); Ubiquone - Q (blue van der Waals representation). Inset: The conformation of the Lys/Glu ion pair (here Lys-204M/Glu-123M) can modulate the pKa of the middle Lys (Lys-235M). (b) The NuoM antiporter-like subunit with water molecules (red dots). The arrow shows the flow of protons through a hydrogen bond-linked network of waters that "visit" the structure during μs molecular dynamics simulation. (c) Snapshot of structures obtained from molecular dynamic simulations of open (in blue) and closed (in red) proton channels from the N-side in Nqo13 (NuoM/ND4), showing conformational changes in the broken helix element. (d ) The structural symmetry of the antiporter-like subunits with an N-side input channel near broken helix TM7b and output channel near broken helix TM12b. Kaila VRI. 2018 Long-range proton-coupled electron transfer in biological energy conversion: towards mechanistic understanding of respiratory complex I. J. R. Soc. Interface 15: 20170916. http://dx.doi.org/10.1098/rsif.2017.0916. Creative Commons Attribution License http://creativecommons.org/licenses/by/4.0/, How is the reduction of ubiquinone and proton transport coupled? The mechanism likely involves conformational changes in flexible regions in the interface of the hydrophilic and membrane domains of the complex. Inhibition of Complex I Complex I is inhibited by more than 60 different families of compounds. They include the classic Complex I inhibitor rotenone and many other synthetic insecticides/acaricides. The classes include Class I/A (the prototype of which is Piericidin A), Class II/B (the prototype of which is Rotenone), and Class C (the prototype of which is Capsaicin). They appear to bind at the same site. Figure $16$ shows their structures. From the structure of the 3 prototypes, what are the characteristics of the pharmacophore, the “ideal binding ligand”? Where do they likely bind? How “promiscuous” is the binding site? Many devastating neurological diseases are associated with defects in Complex I. In addition to major problems with oxidative ATP production, reactive oxygen species (ROS) increase. The major sites for the generation of ROS are Complex I and Complex III. Given the locations of the electron carriers at the periphery and internal within the protein complex, which electron carriers might most readily leak electrons to dioxygen? What ROS is likely to form in the process? Inhibitors might block access to UQ or conformational changes necessary for the final reduction of the ubiquinone free radical. Class A inhibitors dramatically increase ROS production. The actual site of ROS production in Complex I is a bit controversial. One possible electron donor to dioxygen is FMN since both can engage in 1 e- transfers. Mutants that lack N2 iron-sulfur cluster showed ROS production. In submitochondrial preparations, normal Complex I activity occurs (which leads to the formation of a sustained proton gradient). Also reverse electron transport, powered by an artificial proton gradient can occur, which leads to the reduction of NAD+, as shown in Figure $17$. Figure $17$: Overview of forward and reverse electron transport in Complex I. A summary of the finding on superoxide production by Complex I show that: • superoxide production is inhibited by flavin site inhibitors but not Q site inhibitors. • reverse electron transport leads to NAD+ and O2 reduction • Reverse electron transport superoxide production is inhibited by both flavin and Q site inhibitors Figure $18$ shows an expanded view of electron transport and sites where ROS production are enhanced. Refer to the legend for details. Complex III Complex III is a complicated, multisubunit protein that is at the heart of the respirasome. The subunits involved in electron transfer are cytochrome b, cytochrome c1, and the Rieske iron-sulfur protein (ISP). Cytochrome b has two hemes. One is cyto b562 which is also called the low potential heme or cyto bL. The other is cyto b566 which is called the high-potential heme or cyto bH. The cytochrome c1 subunit has one heme. The Rieske iron-sulfur protein contains a Fe2S2 cluster. Figure $19$ shows the relative position of the bound mobile electron carrier, cytochrome C, and the internal ones, the Rieske Fe/S cluster and cytochrome bL and bH. Note also the molecule stigmatellin A, which binds to the site where UQ becomes reduced (called the Qo site) and inhibits the complex. This shows that UQ/UQH2 are in a position to react readily with the Rieske center and cytochrome bL heme. Figure $20$ shows an interactive iCn3D model of a eukaryotic respiratory complex III (3CX5) Figure $20$: eukaryotic respiratory complex III (3CX5). (Copyright; author via source). Click the image for a popup or use this external link:https://structure.ncbi.nlm.nih.gov/i...5riT4GSSGhCHfA. (loads slowly given the size of the structures). The Rieske iron-sulfur protein has a Fe2S2 iron-sulfur cluster which differs from other such clusters in that each Fe is also coordinated to two His side changes, as shown in Figure $21$. Alterations in H bonds to the histidines and to the sulfurs in the complex can dramatically affect the standard reduction potential of the cluster. As with Complexes I and IV, proton and electron transfer are coupled processes. However, in contrast to Complex I, in which protons pass through protein domains that have homology to K+/H+ antiporters, and Complex IV, in which they pass through a combination of a water channel and the H-bond network, the protons in Complex III are carried across the intermembrane space by ubiquinone itself. Two reduced ubiquinones (UQH2) from complex I pass their four matrix-derived protons into the intermembrane space. In the process, four electrons are removed in a multiple-step process called the Q cycle. The two electrons from each UQH2 take different paths. One electron moves to a Fe/S Rieske cluster and the other to cytochrome bL. The electrons moved to the Rieske center then move to cytochrome c1 and then to the mobile electron carrier cytochrome C which is bound to the complex in the intermembrane space. The electrons moved to cyto bL are transferred to cytochrome bH in the complex. Through this latter path, two electrons (from two UQH2) are then moved to oxidized UQ, and two matrix protons are added to reform one UQH2. Hence, only one UQH2 participates in the net reaction shown below. QH2 + 2 cyto c3+ + 2H+matrix → Q + 2 cyto c 2+ + 4H+IMS This net overall reaction, the Q cycle, is illustrated below. This net overall reaction, the Q cycle, is illustrated in Figure $22$. Once again, there are no “proton” channels or hydrogen-bonded networks in the protein for proton transfer across the inner membrane. Another way to think about the electron transfer process from UQH2 to cytochrome C is that the 2 electrons from UQH2 take two different paths, one a high potential path to the Rieske center and on to cytochrome C, and another low potential path to the bL heme and on to the bH heme and then to UQ to reform UQH2 (see figure above). Complex III, along with Complex I, can also produce unwanted reactive oxygen species (ROS). Only three of the protein subunits, cytochrome b (with the bL and bH hemes), cytochrome c1, and the Rieske iron-sulfur protein (ISP) are involved in electron transfer, so one of those is most likely involved in ROS production. Experiments and mathematical models support a mechanism that involves a reduction of UQ by the addition of one electron from cytochrome bL to form UQ. which then passes its electron to dioxygen to form superoxide (O2-.). As two ubiquinones must bind to the complex, there must be two proximal sites. One is the Qi site where oxidized UQ binds and receives an electron. The other is the Qo site where UQH2 binds. From a kinetic perspective, the first UQH2 binds and transfers two electrons, one to the Rieske cluster (and on to cytochrome c1 and then to cytochrome C) and one to cytochrome bL (and on to heme bH) and then to an oxidized UQ bound at the Qi site. The UQ. radical is stabilized by the adjacent bH heme which has a lower affinity for electrons. Now a second UQH2 binds to the Qo site, and transfers two electrons, again one via the Rieske cluster and the second through cytochrome bL and bH to the UQ. radical present at the Qi site to form UQH2 after two protons are transferred to it from the matrix. Now a second UQH2 binds to the Qo site, and transfers two electrons, again one via the Rieske cluster and the second through cytochrome bL and bH to the UQ. radical present at the Qi site to form UQH2 after two protons are transferred to it from the matrix. Antimycin A, an extremely toxic drug, binds to the UQ Qi site and hence blocks electron transfer from cytochrome bL to bH at the Qi site. Heme bL can then pass its electron to dioxygen to produce superoxide. Before we leave Complex III, just a quick observation.  The complex has two identical cytochrome b subunits, each with a bL and bH heme.  The standard reduction potential of the bL and bH hemes in a subunit differ.  This is another example that shows that the standard reduction potential, E0, depends on the environment of the electron acceptor/donor, just as the pKa of an acid depends on its environment. Figure $23$ shows an interactive iCn3D model of just the cytochrome b subunits and all the hemes in the eukaryotic respiratory complex III (3CX5) tp give more clarity to the differing environments of the hemes in the two identical cytochrome b subunits. Figure $20$: Cytochrome b subunits and all the hemes in the eukaryotic respiratory complex III (3CX5). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...BaLQH2KwahNFt9 https://structure.ncbi.nlm.nih.gov/i...BaLQH2KwahNFt9 Complex IV - Cytochrome C oxidase (CCOx) This is the final complex in the electron transport chains and is the one that passes 4 electrons and 4 "substrate" protons to dioxygen to produce water. In addition, it moves as many as 4 H+s from the matrix to the intermembrane space, for a net change of 8 H+s. Here is a modified version of Equation 1 that represents the transported Hs. $\ce{O_2 + 4e^{-} + 4H^{+} + 4H^{+}_(in) → 2H_2O + 4H^{+}_(out)}. \nonumber$ The structure of complex IV is shown in Figure $24$. KEGG pathways (with permission). The 4 H+s that are moved across the membrane are not transported by protein domains that are structural equivalents of antiporters, as in Complex I. Rather elaborate H-bonded water channels with contribution from a space of polar backbone and side chain H-bond donors and acceptors are involved in proton transport. Hence a key question to understand is how electron transfer is coupled to H+ transfer through this hydrogen-bonded network. Since it's quite complicated we will present multiple different but somewhat redundant figures of low to high resolution to help explain this coupled process. Figure $25$ shows an interactive iCn3D model of the 14-subunit human cytochrome c oxidase (5z62) with bound cofactors required for electron transfer. Note that it does not show cytochrome C. Figure $25$: 14-subunit human cytochrome c oxidase (5z62). (Copyright; author via source). Click the image for a popup or use this external link:https://structure.ncbi.nlm.nih.gov/i...XoHRyCkpeRN917. (loads slowly given the size of the structure). Reduced cytochrome C, a mobile electron carrier protein, binds to Complex IV in the intermembrane spaces and singly passes 4 electrons to a series of electron carriers (oxidizing agents) in the protein, eventually to dioxygen as the terminal electron acceptor. Figure $26$ shows the arrangement of the electron acceptor cofactors in Complex IV. Electron transfer presumably occurs through quantum mechanical tunneling. Cytochrome C, the initial “substrate” of this complex, delivers electrons from its heme cofactor to a dinuclear copper cluster, Cu A, where the copper ions are collectively coordinated to two histidines and two cysteines, and a methionine. Figure $27$ shows an interactive iCn3D model of the coordinating ligands for Cu A cluster in Complex IV (5z62) Figure $27$: coordinating ligands for Cu A cluster in Complex IV (5z62). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...7ARphDU4oyDS4A From there, electrons flow to an adjacent heme a (low spin), which has two coordinating histidine ligands. This heme hence does not bind dioxygen. From there, electrons move to another heme, heme a3 (high spin), and then finally to dioxygen which is coordinated to the Fe in heme a3 and to an adjacent Cu B. Heme a3 has only one coordinating histidine ligand, which allows dioxygen binding to the unligated site. The heme a3 Fe:Cu dinuclear cluster is unique among all hemes. Figure $28$ shows an interactive iCn3D model of the coordinating ligands for the hemes and Cu B in Complex IV (5z62) Figure $28$: coordinating ligands for CuB, heme a and heme a3 in Complex IV (5z62). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...4EUooXkwh13Fm8 First, let’s consider the transfer of electrons from heme a to heme a3 and then on to dioxygen (we will consider the entry of electrons into the complex later). If dioxygen, a substrate for the reaction, dissociated from the heme a3 Fe before it was completely reduced, toxic ROS would result. This suggests a reason for the evolution of this key enzyme to have produced the unique heme a3 Fe:Cu dinuclear cluster Heme a and a3 vary from the heme in hemoglobin as they both have a formyl group replacing a methyl and a hydroxyethylfarnesyl group added to a vinyl substituent. Its structure is shown in Figure $29$. What is its overall charge of the heme in its reduced state? In its oxidized state? As hemes are prosthetic groups, they must bind to the apoprotein. The hydroxyethylfarnesyl group, with its nonpolar tail, facilitates the binding of the heme to both the Fe+2 and Fe+3 forms by about 6.3 kcal/mol (26 kJ/mol). The formyl group in heme a increases the E0 value (i.e. make the heme a better oxidizing agent) by about 179 mV (4.1 kcal/mol, 17 kJ/mol). The effect appears to be mediated by the binding of the heme group to the protein again, only in this case there is a differential effect. The formyl group preferentially stabilizes the binding of the Fe+2 heme compared to the Fe+3 heme, which in a thermodynamic cycle, would promote the binding of an electron to Fe+3-heme (increasing its E0 value) to form the more stably bound Fe+2 heme. The key challenge has been to understand the redox coupling to H+ transport. How is this done? The electrostatic environment of the hemes must be considered. In heme a, the two axial ligands are uncharged imidazole side chains of histidines. The overall charge on Fe2+-heme is zero as the two propionates cancel the charges on Fe2+. However, there is a net +1 charge in the Fe3+-heme. This charge could be delocalized within the conjugated pi electrons of the planar heme, which could contribute to the deprotonation of a side chain near heme a in a process that would couple redox and H+ movement. The formyl group of heme a is coplanar with the heme ring in both Fe3+-heme and Fe2+-heme, which allows it to participate in the pi-conjugated electron system of the ring. As the formyl group forms an H bond with Arg 38, changes in oxidation might affect Arg 38 as well. Many other amino acids are involved as well. One is Asp-51 of subunit I which contains heme a. This undergoes a conformational change which moves it to the surface of the inner membrane on reduction. It is near the matrix in the oxidized state. Hence it is likely involved in a proton transport pathway. In fact, crystal structures of oxidized and reduced Complex IV show water channels and small “cavities” which calculations show can hold 1-3 water molecules. Hence groups around the heme, including R38 are assessable to water. Water would be involved in H+ transport through "handshaking" transfer of protons through a hydrogen-bonded network of waters and selective side chains. Some of the amino acid residues associated with the water channels are shown in Figure $30$: and include R38 (the one hydrogen bonded to the formyl group of the heme), S34, T 424, S461, S382, and H413. Another consequence of electron transfer to heme a involves the interaction of S382 and the farnesyl OH group, which are close in space and proximal to a water cavity. On reduction of the heme, a conformation change occurs which increases the standard reduction potential of the S382-farnesyl OH group. What effect would this have on the interaction of the two and the S382-L381-Val380 localized conformation? A new water cavity appears to emerge on reduction in this region. How might this impact proton transfer from the matrix? Now let’s consider the entry site of electrons into the complex and how they might influence proton transport. As mentioned above, Asp 51 (D51) appears to play a key role. It is shown in Figure $29$ above and also in Figure $30$ below. In the oxidized state, D51 interacts with two OH side chains (S205 and S441) and amide NH backbone groups but is not exposed to water. On reduction, D51 lies on the surface in an aqueous environment. Near D51 is Y440-S441. The backbone carbonyl group between 440 and 441 forms an “indirect” interaction with R38 (right panel in Figure $31$), which we showed earlier is affected by the redox state of heme a. They are too far apart to form H bonds. Add two water molecules and envision a bridging interaction between the carbonyl O and the side chain R38 via Y371. Likewise in your mind add a water to allow a bridged hydrogen bond interaction between Y371 also forms a H bond and the heme a propionate. A more nuanced understanding of the mechanism and linkage between H+ and e- movement derives from high-resolution structures determined by Yano et al (2016). In their model (shown in the figures below based on the oxidized form of the protein, pdb 5b1a), protons from the negative (matrix) N side of the complex enter through a water channel and proceed to the positive (intermembrane side) through a H bond network (as described above and depicted below). These comprise the H-Pathway for H+ transfer across the membrane. Two other H+ transfer pathways, the D- and K-pathways, are used as a source of substrate H+ for the 4 H+s added to dioxygen to form 2H2Os. Directional movement is mediated by proton:proton repulsion aided by an increase in + charge on heme a when it transfers an electron to heme a3. Of course, proton:proton repulsion would move protons in both directions. Reverse flow back through the water channel is prevented by a conformational change on oxygen binding that closes the channel. Ultimately 4 electrons are transferred from cytochrome Cs (in single electron steps) to the dicopper cluster, CuA, and then sequentially to heme a to heme a3 (near the copper B ion) to dioxygen to form water. The motion of electrons and protons is coupled electrostatically. Figure $32$ gives an overview of these movements. The small red dots are the oxygen atoms of internal water molecules (the rest have been removed using Pymol). It should be apparent, given the number and location of the internal water molecules, that many would be involved in the proton translocation pathways. What's so interesting about this model is the detailed description of two types of protons, the ones that add to dioxygen and end up in water (substrate protons), and those that are vectorially (directionally) transported to the IMS. In their model, the H+s that end up being transported move through the water and H bond network through a connecting H bond link region to a Mg2+/water cluster. Since the binding of oxygen leads to structural changes that close off the water channel, all protons to be transported to the IMS must be bound in the cluster before dioxygen binding. Figure $33$ shows that initially, 4 H+s move through the H system to the Mg2+/H2O cluster. Oxygen binding then closes the water channels. This buildup of positive charges would certainly lead to enhanced electrostatic attractions for the next phase of the reaction, the movement of electrons into the heme cofactors. Additionally, the 4 H+s in the cluster are probably prevented from leaking to the P side through waters that are proximal (see above figure) by a proline cluster, which presumably restricts the dynamical motion of the protein in that region necessary for proton movement. The figure does not show charge changes in the electron carriers. The figure below breaks downs the mechanism to show the addition of the first electron to the CuA (dicopper cluster), delivered from cytochrome C, and the subsequent transport of one proton from the fully proton-loaded Mg2+/water cluster after dioxygen binding. This figure does show the stepwise redox changes in the electron carriers. After CuA receives an electron from cytochrome C, it donates it to heme a and not to heme a3, even though both are close. The extra negative on heme a facilitates proton pumping though the H pathways shown. Figure $34$s shows a cartoon description of the movement of protons and electrons through Complex IV. Figure $34$: A schematic representation of the role of the Mg/H2O cluster in the proton-pumping mechanism of bovine heart CcO. Yano N et al. J Biol Chem. 2016 Nov 11;291(46):23882-23894. doi: 10.1074/jbc.M115.711770. Epub 2016 Sep 7. PMID: 27605664; PMCID: PMC5104913. Creative Commons Attribution (CC BY 4.0) The Mg/H2O cluster and the H-pathway are connected with the short hydrogen bond network, as labeled in A (a). The location of heme a is shown by Fea3+ or Fea2+ attached to the hydrogen bond network of the H-pathway. The CuA site is on the water-accessible surface of the Mg/H2O cluster. The conformation of the CuA-Mg2+ complex is shown by the shape of a line connecting the two metal ions. The proton-accepting sites are shown by the four hollows on the water-accessible surface of the Mg/H2O cluster. The O2 reduction site is shown by CuB1+ and Fea32+. The pumping and chemical (water-forming) protons are labeled in green and blue, respectively. Electron transfers are shown by red curves. A, overall catalytic cycle of CcO. B, a typical single electron transfer from CuA to the O2 reduction site, coupled with the uptake and release of protons. For the sake of simplicity, the uptake of one chemical proton equivalent (the average number) is given in B. Various oxidation and ligand binding states in both CuB and Fea3 in the intermediate states shown by A (d) and B (d) are not included for the sake of simplicity. Figure $35$ converts the schematic representation into a "structural cartoon" representation showing one cycle of coupled electron and proton transfer into Complex IV. The H- (for transport of 4H+s across the membrane) and the D- and K-pathways (sources of substrate 4H+s for water synthesis from O2 are shown in Figure $36$. Two black arrows show the D- and K- for substrate H+s that form water, while the blue arrow shows the H-pathway for the hydrogen bonding network of water in the water channel. The Mg/H2O cluster (blue area) is attached to the hydrogen-bond network of the H-pathway by a short hydrogen-bond network (gray area with water shown as blue sphere). The formyl group and one of the propionate groups of heme a are hydrogen-bonded with Arg38 and a fixed water molecule in the hydrogen-bond network of the H-pathway. Compare this diagram to the ones presented in the figures above it. Again we show multiple representations to give readers many opportunities to conceptualize this complicated complex. Complex II - succinate:ubiquinone oxidoreductase (SQR) - succinate dehydrogenase You've seen complex II before with another name, succinate dehydrogenase from step 6 of the citric acid cycle, which catalyzes the following reaction. succinate + FAD ↔ fumarate + FADH2 ΔGo = 0 kcal/mol The ΔGo for this reaction is about 0 kcal/mol so it is readily reversible. However, something vital is left out of this description. The above reaction appears to suggest that the FAD/FADH2 are readily diffusable and bind and unbind after catalysis. That's not true. Instead, FAD/FADH2 are covalently attached to the enzyme so after one cycle of enzyme catalysis, the enzyme is dead. Another substrate/product pair must interact with enzymes to reconvert the covalently attached FADH2 back to FAD so catalysis can continue. The other pair is ubiquinone (UQ/UQH2). A better description of the reaction equation is shown in Figure $37$. Complex II clearly links the tricarboxylic acid cycle to electron transport! Figure $37$: Complex II - Succinate Dehydrogenase Reaction As with the other mitochondrial electron transport complexes, Complex II is a multisubunit integral membrane protein. The hydrophilic section protruding into the matrix contains two proteins, a flavin-binding protein (Fp) and an iron/sulfur protein (Ip). The transmembrane domain has two "cytochrome binding proteins", a lighter one (CybL) and a heavier one (CybS). The complex has binding sites for dicarboxylic acids (succinate/fumarate) and two binding sites for ubiquinone, a proximal higher affinity site (Qp) on the matrix side closer to the succinate binding site, and a distal lower affinity site (Qd) near. The ubiquinone sites bind 2-thenoyltrifluoroacetone (TTFA) which acts as an inhibitor of ubiquinone reduction. 3-nitropropionate - (NO2)(CH2)2CO2- is structural similar to succinate - (CO2-)(CH2)2CO2- and is an inhibitor of succinate oxidation. Figure $38$ shows an interactive iCn3D model of the Mitochondrial Respiratory Complex II from porcine (pig) heart (1ZOY). The chains are: • hydrophilic FAD-binding protein (lavender) • hydrophilic FeS protein (blue) • integral membrane large cytochrome binding protein (brown) • integral membrane small cytochrome binding protein (green) Figure $38$: Mitochondrial Respiratory Complex II from porcine heart (1ZOY) . (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?KKPWG9dW1ARq3wQf9. (loads slowly given the size of the structure). Given the role of the enzyme in both the mitochondrial citric acid and electron transfer pathways, you would expect high electron transfer rates between the dicarboxylic acid and the ubiquinone binding sites. Any disruptions in electron transfer (by inhibitors or mutations) could lead to the production of toxic reactive oxygen species (ROS). Binding an inhibitor to the proximal Qp site would lead to a buildup of FADH2 which could transfer electrons in single-electron steps to dioxygen to produce ROS. This would occur if the dicarboxylic site were empty. As there is no H+ transfer from the matrix to the intermembrane space for Complex II, we'll only focus on electron transfer from succinate to FAD and on to ubiquinone. The transfer of electrons from succinate to ubiquinone hence takes place in two steps. Step 1: Succinate + FAD ↔ Fumarate + FADH2. Figure $39$ shows a plausible mechanism using a histidine in the active site as a general base. This is a simple hydride (2e-) transfer reaction to FAD. Note that no FeS cofactors are required. Step 2: FADH2 + UQ ↔ FAD + UQH2. This does not proceed by transfer of 2 e- from FADH2 directly to UQ. Rather it occurs through a series of the FeS cofactors (Fe2S2, Fe4S4, and Fe3S4 clusters) and heme. First, let's consider the path of electron transfer which will occur in single electron steps through the intermediary FeS clusters to ubiquinone as shown in Figure $40$. The standard reduction potential E0 is shown for each step. Note that FAD is shown instead of the actual electron donor, FADH2, to correspond to the standard reduction E0 values shown in tables. Electrons flow from FADH2 (lowest, most negative E0 value, -79 mV) to UQ (highest, most positive value E0 value, +113 mV) or from weakest to strongest oxidizing agent. Electron transfer depends on distance as well. It appears that the transfer of electrons to heme b is not likely, given the presence of the closer UQ with the most positive E0 value. Hence the function of heme b is unclear. However, if electrons are transferred to it, it would quickly and favorably send them on to UQ. Note that the FeS clusters have progressively more positive E0 values except for Fe4S4 whose E0 = -260 mV. Its E0 is depressed by the presence of charged/polar groups which would make the transfer of a negative electron less likely. The net transfer standard potential is clearly favored thermodynamically. As the E0 and corresponding G0 values are state functions, the final ΔG0 doesn't depend on the path. The high value of E0 for Fe4S4 presents an activation energy barrier for transfer in both directions and probably prevents reverse electron transport in this complex. The membrane domains have two polar (Asn, Ser) and 5 polar-charged (Asp, Glu, and Lys) side chains. Now let's look at the terminal step in the transfer of electrons from Fe3S4 to ubiquinone. Figure $41$ shows a plausible mechanism. Ubiquinone is shown in black. TTFA, which binds to the ubiquinone site and inhibits electron transfer, is shown in brackets. Tyrosine 91, Serine 41, and Histidine 216 are likely candidates for involvement in catalysis as shown below. Arginine 46 and aspartate 90 might be involved in the protonation steps of UQH2. Figure $42$ shows an interactive iCn3D model of the iCn3D TTFA1 binding site (where ubiquinone binds). Figure $42$: Mitochondrial Respiratory Complex II bound with 2-thenoyltrifluoroacetone (1zp0). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?dY3niSfLVa9WiNNe7

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/19%3A_Oxidative_Phosphorylation/19.01%3A_Electron-Transfer_Reactions_in_Mitochondria.txt

Search Fundamentals of Biochemistry Introduction ATP synthase, also called F1FoATPase, is a rotary motor enzyme. This enzyme is found in the inner membrane of mitochondria, the analogous thylakoid membranes of chloroplasts, and the cell membrane of bacteria. The enzyme consists of two parts, the membrane-bound Fo which is a proton translocator, and the F1 part which has catalytic (ATP synthesis or hydrolysis activity. The Fo part can be considered to be a rotary electrical motor powered by proton flow, while the F1 part acts as a rotary chemical motor powered (in reverse) by ATP hydrolysis. The Fpart is named since it is sensitive to oligomycin (note that it should theoretically be read as Fo and not Fzero). The F1 part is named since it was eluted from a column chromatography column in Fraction 1. The elucidation of its structure and mechanism by many but especially by Paul D. Boyer and John E. Walker, was a major feat of scientific study. They, along with Jen Skou, who studied the mechanism of the analogous Na/K ATPase, were awarded the Nobel prize for their work. Let's start with the known structure of this complicated membrane-bound rotary enzyme and then work towards an understanding of how it works. The structure of the bovine F1FoATPase is shown in Figure $1$. Figure $1$: Bovine F1FoATPase. In the previous section, we discussed how oxidative electron transport in the mitochondria is accompanied by proton transport against a concentration gradient from the matrix through the inner member to the intermembrane space. The synthesis of ATP from ADP and P1 is endergonic and requires an energy source. That energy source is provided by the thermodynamically favored collapse of the pH gradient across the mitochondrial inner membrane. Protons flow through the Fo membrane spanning C helices from the intermembrane space (not shown in Figure 1) to the matrix. This powers the synthesis of ATP from ADP and Pi by the F1 part of the enzyme in the matrix lumen. Figure $2$ shows an interactive iCn3D model of the entire bovine mitochondrial ATP synthase (5ARA). The subunits are shown in the same color as Figure 1 above. Figure $2$: Bovine mitochondrial ATP synthase (5ARA). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...3YUBWrX91nnua9 The enzyme is reversible. If protons flow down a concentration gradient through Fo, ATP is synthesized by F1. Alternatively, ATP hydrolysis by F1 leads to the transport of protons through Fo and against a concentration gradient. Isolated F1 can only break down ATP, and not synthesize it. We called this structure a rotary enzyme, so what rotates? A rotary "axle" protein complex comprised of the γ, ε, and δ subunits (some also describe the γ, ε, and δ subunits as part of F1) rotates. Its interactions with the multimeric complex of c alpha helices (shown in purple in the figures above) in the main Fo complex causes the "c-ring complex" complex in Fo to rotate as well. The F1 part does not rotate because of the conformational stability of the β subunit and the connection to the long alpha helices of the D and B1 proteins, which comprise the "stator (the stationary) part of an electric motor", which keeps F1 stationary. Hence the enzyme does truly act like an electric motor, whose components are shown in Figure $3$. Figure $3$: Electric motor with rotor, shaft, and stator. http://www.cdmmotor.com/en/article-22982-28485.html Now let's look at the mechanisms and the evidence for them for each part of this remarkable enzyme. F1 The F1 unit (with a quaternary structure of a3β3 forming a hexagonal ringed structure with a central cavity, occupied by a gamma subunit)is about 80 angstroms from the Fo subunit and both are connected to the rod-shaped γ subunit which spans the center of the a3β3 ring. Energy transduction (necessary to capture the negative free energy change associated with the collapse of the proton gradient to drive the positive free energy change for ATP synthesis) occurs between the two subunits. Figure $4$ shows a top view of the α (red) and β (green) subunits in the a3β3 F1 complex Figure $4$: Top view of the α (red) and β (green) subunits in the a3β3 The γ subunit is shown coming out of the plane of the ring. ATP is shown in gold spheres, ADP in purple, and the smaller SO42- (from a crystallizing agent) is shown in CPK spheres. They are sandwiched between the a and β subunits. Boyer, in the absence of the complete structure of the F1Fo ATP synthase, was able to deduce from experimental evidence that the a3β3 complex, which can be viewed as three aβ dimers (with catalysis occurring between subunits of individual dimers where ATP and ADP bind), have three different, interconvertible conformation defined as a Loose (L), Open (O) and Tight (T) states, with names describing the strength of substrate binding in each dimer. • O - open state with very low affinity for substrates and has no catalytic activity; • L - loose state with low affinity for substrates and also no catalytic activity; • T - tight state with high affinity for substrates and with catalytic activity. The three states rotate not physically with respect to some central axis but conformationally, depending on their interaction with the γ subunit which binds perpendicularly in the central junction of the a3β3 ring. Changes in the orientation of the central γ subunit due to its rotation with respect to the a3β3 ring cause the conformation of the O, L, and T states to change in situ with the orientation of the rotating γ subunit The conversion of the LOT conformations, their binding of substrates (ADP and Pi), the conversion of bound ADP and Pi, and the release of the product (ATP) proposed by Boyer are shown in Figure $5$. Figure $5$: Boyer's three-state conformational model (L-O-T) for ATP synthesis The collapse of the proton gradient (i.e. the proton-motive force) causes the γ subunit to rotate like a crankshaft relative to the F1 subunit, forcing the β subunit to change conformation from the T to the O (releasing ATP) and then to the L (binding ADP and Pi) states. The γ subunit does not appear to undergo any significant conformational change on ATP hydrolysis as evidenced by tritium exchange studies of amide protons. To prove that the γ subunit rotates, you'd have to observe a single molecule. Since the γ subunit was too small to visually discern its rotation, Noji et al covalently attached a fluorescein-labeled actin filament to the γ subunit (near where Fo would bind). The whole F1 molecule was fixed to a glass slip through a His-tag such that the a3β3 ring was effectively immobilized. The γ subunit was free to rotate, which could be detected by observing the fluorescence under a fluorescent microscope from the attached actin filament. This experiment and the outcomes are described in Figure $6$. Figure $6$: The direct observation of the γ rotation in the F1 motor. Noji and Yoshida. JBC (2001) DOI:https://doi.org/10.1074/jbc.R000021200. Creative Commons Attribution (CC BY 4.0) Panel A shows the experimental system for the observation of the γ rotation using an optical microscope. The F1 motor tagged with 10 His residues at the N terminus of the β subunit was immobilized upside down on a coverslip coated with nickel-nitrilotriacetic acid (Ni-NTA). An actin filament (green) labeled with fluorescent dyes and biotins was attached to the biotinylated γ subunit (gray) through streptavidin (blue). Panel B shows the rotary movement of an actin filament observed from the bottom, the membrane side, with an epifluorescent microscope. Length from the axis to tip, 2.6 μm; rotary rate, 0.5 revolutions per s; the time interval between images, 133 ms. The actin filament rotated only in the presence of ATP. It rotated only counterclockwise, indicating that the motion was not random, but a specific motion of the γ subunit. At extremely low concentrations of ATP, rotation occurred only in 120o increments, implying one step per molecule of ATP hydrolyzed. (Remember the β subunits are separated by 120o ). As the rotation occurs, there is viscous resistance to the movement of the actin filament. He calculated that for a single 120o step caused by hydrolysis of a single ATP molecule, the amount of work was 80 piconewton which is about the free energy of hydrolysis of a single ATP molecule. Later experiments in which a colloidal gold nanoparticle (40 nm diameter, with less frictional resistance to movement) was used instead of an actin filament showed the same result. At low [ATP], the motor rotates in 120o steps. At high [ATP], the rotation rate becomes continuous and saturates (with Michaelis/Menten kinetics) at 130 revolutions per second. Other experiments using immobilized ATPase and magnetic tweezers have addressed the timing of substrate binding and product release when the enzyme is run in reverse (ATP hydrolysis). On rotation of the γ subunit, the three binding sites change properties. In hydrolysis, ATP binds to the open site and helps promote the 120-degree rotation. In the next step, ATP is hydrolyzed. In the final step, products dissociate. Pi dissociation occurs last from the third site. Hence each of the 3 beta-binding sites has different roles. One binds the substrate, one performs catalysis and third releases products. Assuming the synthesis pathway is the reverse of the ATPase reaction, the final release of Pi in ATP cleavage predicts that Pi binds first in the synthetic direction. This would preclude the binding of ATP next which is critical since its concentration during synthesis can be 10x higher than that of ADP. As Pi is bound first, only ADP, not ATP can bind next. The γ subunit rotation plays a "catalytic" role as its rotation induces cyclic conformational changes in the beta subunit of the synthase. Can ATP synthesis occur without the gamma subunit by a mechanism that involves a less proficient, but a concerted set of cyclic changes in beta subunit conformation? It can. Uchihashi et al have used high-speed atomic force microscopy (AFM) to study the a3β3 ring from the F1 subunit without the gamma subunit. They found that upon ATP hydrolysis, the beta subunits underwent conformational changes in the same counterclockwise rotary direction as when the gamma subunit was present. This is illustrated in Figure $7$. Figure $7$: AFM Study of Conformational Changes in F1 "gammaless" subunit These experiments conclusively show that the F1 subunit is effectively a rotary motor with the gamma subunit acting as a rotor in the stationary hexagon ring composed of the 3 pairs of alpha/beta subunits which acts as the stator (stationary part of an electric rotary motor). The actual amino acids involved in the mechanism of ATP synthesis/hydrolysis are still not clearly defined but Glu 190 on the beta subunit acts as a general base. Figure $8$ shows bound ADP and the proximity of Glu 188. Ala 158 is thought to move towards the active site after a conformational change, with the nonpolar methyl side chain displacing an adjacent water molecule which could leave as a product of ATP synthesis. Figure $8$: Active site residues for the synthesis of ATP. ADP is shown in spacefill, and CPK colors. Mg2+ is shown as a gray sphere (1E79). Fo and Proton Transfer The mechanism by which the proton gradient drives ATP synthesis involves a complex coupling of the Fo and F1 subunits. Proton translocation occurs through the interface of the inner mitochondrial (or cell membrane of bacteria) membrane proteins C1 and A (or beta), whose structures from the bovine enzyme are shown again in Figure $8$. Figure $8$: Inner membrane proteins C1 and A (or beta) For protons to translocate through the membrane, there must be an aqueous channel on the matrix side and the lumenal side (bounded by the inner and outer mitochondrial membranes). So we have two understand two features: • how the rotary axle part of the complex interacts with the C1 hexameric ring, causing the ring to rotate in the membrane. • The nature of the water channels and the pathway for proton translocation. The number of c monomers in the c-ring can vary between 8-17. For example, the bovine ATP synthase shown in Figures 1 and 2 has 8 c-monomers. Each has a critical proton donor and acceptor located near the a or beta-Fo chain. In the bovine case, it is glutamate 58. (The side chains in the bovine structure presented in Figure 2 do not show since they were not resolved in the cryoEM structure.) In E. Coli, which we will explore further below, it is aspartate 61. In yeast, it is at position 59. They are positioned near the center of the membrane helices. Mutations in these residues cause significant decreases in enzyme activity. An E56Q mutation in some bacilli species prevents proton pumping and ATP synthesis. Another key charged group, arginine, in the a or beta Fo protein is also key. Figure $9$ shows an interactive iCn3D model of the E. Coli A1-C12 subcomplex of F1FO ATP Synthase (1C17). Figure $9$: E. Coli A1-C12 Subcomplex OF F1FO ATP Synthase (1C17). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...sfRJBHEBiC5sy9 The key aspartate 61 (sticks and labeled) in each C ring subunit (red) and the proximity of two of the aspartate 61s to arginine 210 in the A subunit (brown) are evident. Two classic inhibitors (structures shown below) of ATP synthase interact with the Fo subunit. One, oligomycin A, binds between the a and c subunits and blocks proton transport activity of the Fo subunit. The O protein at the top of the F1 complex is also called the Oligomycin-sensitivity-conferring protein (OSCP), even though oligomycin does not bind there. The soluble F1 by itself is not sensitive to oligomycin, but when it's linked to Fo in part through the O or OSCP peripheral stalk protein, it becomes sensitive to oligomycin. Another inhibitor, dicyclohexylcarbodiimide reacts with a protonated Asp 61 in c subunits of Fo. It does so even at pH 8.0 which indicates that the pKa of the Asp 61 is much higher than usual. This might occur if the Asp is in a very hydrophobic environment. The modification of one Asp 61 in only one c subunit is necessary to stop Fo activity. The protonated carboxyl group donates a proton to a nitrogen atom in DCCD, which then reacts with the deprotonated Asp to form an O-acyl isourea derivative. Figure $10$ shows the structures of oligomycin A and DCCD, inhibitors of proton transport by Fo. Figure $10$: structures of oligomycin A and DCCD Figure $11$ shows an interactive iCn3D model of the F1FoATPase c10 ring with bound oligomycin (4F4S) Figure $11$: E. Coli A1-C12 Subcomplex OF F1FO ATP Synthase (1C17). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...pJw1pFr8jTPmK8 Only two of the c-subunits are shown interacting with one oligomycin. The actual complex has 10 c-subunits and 8 oligomycins bound. The oligomycin is shown in spacefill with CPK colors. The two c-subunits are shown in gray. The side chains from the two c-subunits interacting with oligomycin are shown as CPK-colored sticks. They are conserved between humans and yeast. Note the single H bond between the critical Glu 59 and oligomycin. The green colors indicate hydrophobic parts of the c-subunits which interact with the mostly nonpolar parts of the antibiotic. How do these key Asp or Glu side chains participate in proton translocation across the membrane and c-ring rotation and how do they access the matrix and lumenal water channels? We can start to understand the answer to these question by examining the structure of the c-ring and its interaction with the a-chain from Bacillus PS3 ATP synthase, as shown in Figure $12$, which show the key Glu59 residues in each of the c subunits of the 10-mer ring along with Arg 176 and two other glutamates (Glu 223 and Glu 162) in the A ring. Figure $12$: Schematic picture of the a-subunit and c-ring of Fo. Mitome et al. (2022). eLife 11:e69096. https://doi.org/10.7554/eLife.69096. Creative Commons Attribution License Panel (a) shows the ac10 part of the Fo region with the key amino acids shown as colored spheres (and numbered based on the yeast protein). Panel (b) shows a cartoon of a simulation model developed by Mitome et al (ibid). The Glu 59s are shown as green circles and labeled a-j. The gray represents the membrane. Protons move between the protonated Glu 59 in c subunits that are proximal to the deprotonated Glu 223 and Glu 162 in a hydrogen bond "handshake". The Glu 59s in the c chains in the 10-mer ring are likely to be protonated given their position in the center of the membrane, whereas the proximal Glu 223 and 162 are more likely to be initially deprotonated given their proximity to aqueous channels, where they "hand off" their protons to complete the translocation from the intermembrane luminal space to the matrix. Now of course, once a c-ring Glu 59 is deprotonated, it has a negative charge. Simulations suggest that there are 2-3 deprotonated Glu 59 with a negative charge proximal to the a-subunit. One is likely to take up a proton in a process leading to ATP synthesis. The orange arrows show the direction of proton flow. If the c-ring rotates in the clockwise direction, ATP is hydrolyzed while counter-clockwise rotation leads to ATP synthesis. Now let's look at the water channel forming at the interface between the c-ring and a-subunit and an additional protein ASA6 from the unicellular green alga Polytomella sp. The interactions and nature of the water channels are shown in Figure $13$. Figure $13$: Two aqueous channels in Fo. Niklas Klusch, Bonnie J Murphy, Deryck J Mills, Özkan Yildiz, Werner Kühlbrandt (2017) Structural basis of proton translocation and force generation in mitochondrial ATP synthase eLife 6:e33274. https://doi.org/10.7554/eLife.33274. Creative Commons Attribution License Subunit-a is shown in blue, the c10-ring in yellow, and the associated ASA 6 protein in brick. The channels are shown as potential surfaces (red, negative; blue, positive; grey, neutral). (A) and (B) display a 5 Å slice of the c10-ring at the level of the protonated cGlu111. Panel(A) shows a 5 Å slice of the c10-ring at the level of the protonated c-chain Glu111, with the lumenal channel seen from the crista lumen. Panel (B) shows a 5 Å slice of the c10-ring at the level of the protonated c-chain Glu111, with the matrix channel seen from the matrix. Panel (C) shows a side view of both channels seen from the c-ring, with outer c-ring helices in transparent yellow. Lumenal channel, left; matrix channel, right. The strictly conserved a-chain Arg239 in helix 5 (H5) separates the lumenal and matrix channels. Panel (D) shows that the lumenal channel passes through the H5/H6 hairpin at the small sidechains aAla246, aGly247 (H5), and aAla292 (H6) (green). Panel (E) shows H4, the N-terminal half of H5, and the connecting H4/H5 loop at the matrix channel. Figure $14$ a probable proton pathway through the Fo subcomple Figure $14$: Proton pathway through the Fo subcomplex. Klusch et al, ibid. The a-subunit is shown in blue, the adjacent c-ring helices in transparent yellow, and the aqueous channels in translucent grey. Panel (A) shows the movement of protons (red arrow) in the lumenal channel. They likely move between Glu172, His248, His252, and Glu288 in the a-chain (dashed red ellipse) through the H5/H6 helix hairpin at the small sidechains of a-chain Ala246 and Gly247 (H5) and Ala292 (H6) (green) to c-chain Glu111 in the rotor ring c-subunits (red circles). Panel (B) shows that a-chain Arg239 (blue circle) is located halfway between the lumenal channel on the left and the matrix channel on the right, forming a seal to prevent proton leakage. c-ring helices (transparent yellow) with cGlu111 are seen in the foreground. Panel (C) shows the movement of protons (dashed red arrow) in the matrix channel as they pass directly from the deprotonated c-chain Glu111 to the pH 8 matrix. A final summary view showing c-ring rotation powered by the collapse of the pH between the lumenal (intermembrane space) channel (pink) and matrix channel (light blue) is shown in Figure $15$. Figure $15$: c-ring rotation is powered by the potential gradient between the lumenal channel (pink) and matrix channel (light blue). Klusch et al, ibid The c-ring (yellow) and the membrane-intrinsic four-helix bundle of subunit a (blue) are drawn to scale as seen from the matrix. Protons (red) pass from the crista lumen below the projection plane through the lumenal channel between H5 and H6 to protonate Glu111 of c-subunit A, while subunit J is deprotonated by the higher pH of the matrix channel. The positively charged a-chain Arg239 is likely to interact with the deprotonated c-chain Glu111 during its short passage to the lumenal channel. The lumenal and matrix channels approach one another to within 5–7 Å. A protonmotive force of 200 mV between the closely spaced channels creates a local electrostatic field in the range of 40 million to 100 million V/m, depending on the protein dielectric. The field exerts a force on the deprotonated c-chain Glu111 that results in a net counter-clockwise rotation of the c-ring (grey arrow). Scale bar, 10 Å. Click the images below to see (in a new window) a series of stunning videos showing proton translocation across the channel! (Niklas Klusch, Bonnie J Murphy, Deryck J Mills, Özkan Yildiz, Werner Kühlbrandt (2017) Structural basis of proton translocation and force generation in mitochondrial ATP synthase eLife 6:e33274. https://doi.org/10.7554/eLife.33274. Creative Commons Attribution License) The three-dimensional arrangement of subunit a (blue), c-ring (yellow) with the lumenal channel in pink and matrix channel in light blue. Arrangement of channel-lining sidechains for the lumenal channel. Sidechains in stick representation are colored as subunit a, blue; c-ring, yellow; ASA 6, brick; lipids, grey. Channels are shown as potential surfaces (red, negative; blue, positive; grey, neutral). Arrangement of channel-lining sidechains for the matrix channel. Sidechains in stick representation are colored as subunit a, blue; c-ring, yellow; ASA 6, brick. Channels are shown as potential surfaces (red, negative; blue, positive; grey, neutral). In summary, F1FoATP synthase is a rotary enzyme that ultimately couples the collapse of a proton gradient (a chemical potential gradient that contributes to the transmembrane electrical potential) to a chemical (phosphorylation) step. The rotor, which is in contact with both the Fo proton pore and the F1 synthase, moves with respect to both subunits, which couples them. Both the axle and c-ring of Fo rotate. The interaction of the delta protein with the F1 a3β3 hexamer does not cause a3β3 to physically rotate as a whole, as does the c-ring. Rather it causes a sequential rotation in the conformation of the three aβ dimers from the Loose to Open to Tight (LOT) conformations. The Fo pore can hence be considered an electrical motor and the F1 synthase a chemical motor. Carrying the analogy of a motor even further, the Felectrical motor turns the F1 chemical motor into a generator, not of electricity but of ATP. The figure and link below, taken from the Protein Data Bank, go into more depth about this nanomotor. The concerted movement of the c-ring and axle protein with respect to the not physically rotating a3β3 F1 part can be seen in Figure $16$, which is derived from cryoEM structure of different rotated states of bovine ATP synthase. Figure $16$: Structure and conformational states of the bovine mitochondrial ATP synthase (5ARA, 5ARE, 5ARH, 5ARI, 5FIJ, 5FIK, 5FIL) A phenomena video is available showing the detailed steps in ATP synthesis by ATP synthesis Click on Figure $17$ to get the link to the video. Figure $17$: ATP synthase in action. Muzzey and Lue, HHMI Institute. For educational and non-commercial use only. This amazing enzyme can be purified and reconstituted in a lipid vesicle along with bacteriorhodopsin. If these proteins are oriented in the correct fashion, light can cause proton translocation into the lumen of the vesicle creating a transmembrane potential. The proton gradient can collapse as protons move from the lumen through the Fo part of F1Fo ATP synthase, causing ATP synthesis from ADP and Pi in the outside solution. This is shown in Figure $18$. Figure $18$:. Light-driven ATP production. Ahmad et al. ACS Synthetic Biology 2021. https://pubs.acs.org/doi/10.1021/acssynbio.1c00071. https://creativecommons.org/licenses/by/4.0/ Panel(A) shows a cartoon of the light-driven ATP synthesis in lipid vesicles. Panel (B) shows the increase in [ATP] over time. Proton Gradient Collapse and ATP synthesis - Thermodynamics Mathematical analyses show that the free energy change on proton gradient collapse can easily power the endergonic synthesis of ATP. Consider a typical pH gradient (-1.4 pH units) across the inner membrane of respiring mitochondria (with the outside having a lower pH than the inside making the inside more depleted in protons). There is a chemical potential difference in protons across the membrane. However, another factor determines the thermodynamic driving force for proton translocation across the membrane. A transmembrane potential exists across the inner membrane of the mitochondria, as it does across most membranes. The source of the membrane potential will be discussed in the signal transduction chapter (Chapter 28). The inside is more negative than the outside, giving the membrane a transmembrane electrical potential. of about -0.14 V. Clearly, protons would be attracted to the other side of the membrane (into the matrix) by this potential difference, which then augments the chemical potential difference as well. A simple mathematical derivation shows that indeed, a proton gradient can supply enough energy for ATP synthesis, especially when coupled to a transmembrane electrical potential. We reshow a small image of the complex and its activities in Figure $19$ to help you through this section Figure $19$: ATP synthase and its activities. Noji, H., Ueno, H. & Kobayashi, R. Correlation between the numbers of rotation steps in the ATPase and proton-conducting domains of F- and V-ATPases. Biophys Rev 12, 303–307 (2020). https://doi.org/10.1007/s12551-020-00668-7. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/. Chemical Potential (use G instead of μ) Since most students use free energy and not chemical potential (which is just free energy /mole at constant T and P, we will derive these equations using free energy G. Let's consider the free energy of the protons on the outside (low pH, higher concentration) and in the inside (matrix, higher pH, low concentration): \mathrm{H}^{+}{ }_{\text {out }} \longrightarrow \mathrm{H}^{+}{ }_{\text {in }} \mathrm{G}=\mu=\text { chemical potential }(\mathrm{kcal} / \mathrm{mol}) Now express this as the change in free energy, ΔG \Delta G_{\text {chem }}=G_{\text {in }}-G_{\text {out }}=G^{0}+R T \ln \left[H^{+}\right]_{\text {in }}-\left(G^{0}+R T \ln \left[H^{+}\right]_{\text {out }}\right) This can be rewritten as: \Delta G_{\text {ohem }}=\operatorname{RT|n} \frac{\left[H^{+}\right]_{\text {in }}}{\left[H^{+}\right]_{\text {out }}}=2.303 R T\left[-\log \left[H^{+}\right]_{\text {out }}-\left(-\log \left[H^{+}\right]_{\text {in }}\right)\right] Hence \Delta \mathrm{G}_{\text {chem }}=2.303 \mathrm{RT}\left(\mathrm{pH}_{\text {out }}-\mathrm{pH}_{\text {in }}\right)=2.303 \mathrm{RT} \Delta \mathrm{pH} In respiring mitochondria, ΔpH = -1.4, so \Delta \mathrm{G}{ }_{\text {chem }}=2.303 \mathrm{RT} \Delta \mathrm{pH}=2.303(1.99 \mathrm{cal} / \mathrm{mol} \cdot \mathrm{K})(298 \mathrm{~K})(-1.4)=-1.91 \mathrm{kcal} / \mathrm{mol} \mathrm{} \mathrm{H}^{+} Electrical Potential Consider the relationship between ΔG and the transmembrane electrical potential \Delta G_{\text {elect }}=+Z F \Delta \psi where Δψ is the transmembrane potential, Z=+1 (charge on a proton) and F is the Faraday constant This equation is similar to \Delta G \text { elect }=-n F \Delta E where ΔE is the cell potential or EMF. If ΔE is + and Δψ is -, then ΔG is -. In respiring mitochondria, Δψ = -0.140 V (remember that 1 V = 1 J/C). Hence \Delta G_{\text {elect }}=+Z F \Delta \Psi=1(96.485 \mathrm{C} / \mathrm{mol})(-0.140 \mathrm{~J} / \mathrm{C}) \times(1 \mathrm{cal} / 4.18 \mathrm{~J})=-3.23 \mathrm{kcal} / \mathrm{mol} \mathrm{H}{ }^{+} Electrochemical potential Now let's add the equations to get the full driving force, the electrochemical potential: \Delta G_{\text {tot }}=2.303 \mathrm{RT} \Delta \mathrm{pH}+\mathrm{ZF} \Delta \psi=-5.15 \mathrm{kcal} / \mathrm{mol} \mathrm{H}{ }^{+} This gives \frac{\Delta G_{\text {tot }}}{F}=\frac{2.303 R T \Delta p H++z F \Delta \psi}{F} Hence \Delta p=\Delta \psi+\frac{2.303 R T \Delta p H}{F}=\Delta \psi+0.059 \Delta p H=0.224 \mathrm{~V} where Δp is the protonmotive force.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/19%3A_Oxidative_Phosphorylation/19.02%3A_ATP_Synthesis.txt

Search Fundamentals of Biochemistry Introduction The main function of oxidative phosphorylation (oxphos) is to produce, under aerobic conditions, lots of ATP. It should make sense that to a first approximation, the regulation of oxphos depends primarily on the energy state of the cell, which is reflected by the ADP/ATP ratio. In contrast to glycolysis and the citric acid cycle, which produce NADH, the primary and first electron carrier in oxphos, the regulation does not depend on the binding of allosteric effectors to key enzymes in the pathway. Likewise, hormonal regulation is not involved. Perhaps this occurs since the oxphos pathway is downstream from the glycolytic and citric acid pathways, where allosteric and hormonal regulation robustly affects their outputs. If you consider the primary reactants in electron transport to be NADH, FADH2, O2, ADP, and Pi and if you assume that O2 levels are constant, ADP appears to be the main molecule that determines the rate of the oxphos reactions. This is in accordance with what we have previously described: AMP, ADP, and ATP levels regulate many of the key enzymes in the glycolytic and citric acid pathways, as reviewed in Figure \(1\). Figure \(1\): Regulation of glycolysis, the citric acid cycle, and oxphos by ATP, ADP, and AMP. Overall there are several levels of oxphos regulation, which occurs at the level of the kinetics of the electron transport chain complexes, the efficiency of electron and proton transfer, the structure of the mitochondria, and mitochondria formation and their degradation. Coupling and uncoupling of electron and proton transport What is key is that the regulation of oxidation (electron transport) is coupled to the regulation of ATP synthesis (phosphorylation). These processes can also be uncoupled if electrons "leak" away from their transfer from NADH to O2, which leads to the production of harmful reactive oxygen species (ROS) such as superoxide. Proton can also "leak" away from the proton gradient if they bypass the ATP synthase complex and pass through the inner membrane with the help of specific carriers and protein uncouplers, as shown in Figure \(2\). Figure \(2\): Coupling and uncoupling of oxidative phosphorylation. Eyenga, P.; Rey, B.; Eyenga, L.; Sheu, S.-S. Regulation of Oxidative Phosphorylation of Liver Mitochondria in Sepsis. Cells 2022, 11, 1598. https://doi.org/10.3390/cells11101598. Creative Commons Attribution License The far right light green proton and the insert show the proton leak’s kinetics. The dotted line indicates the electron pathways. There are two types of proton leaks: • a basal leak through mitochondrial anion carriers in a non-regulated and minor process. • an inducible leak through specific proteins, adenine nucleotide translocase (ANT), and uncoupling proteins (UCPs), which leads to loss of energy as heat. They can be activated by fatty acids as well as ROS. A particular UCP, UCP1, is found most abundantly in brown fat in mammals. The basal leak occurs in the presence of inhibitors of ANT (carboxyatractylate) and UCP1 (GDP). It can be significant (about 25% of the resting metabolic rate of liver cells and up to 50% in respiring skeletal muscle cells). It is less understood than inducible leaks and does appear to be attributed to ANT in some fashion. When oligomycin, which inhibits the collapse of the proton gradient through ATP synthase, is added to mitochondria, the proton gradient (protonmotive force, Δp) decreases. The rate of leakage increases exponentially with increasing Δp, so it represents non-Ohmic leakage. Ohmic leakage would give a linear dependence on Δp. This can be seen from a rearrangement of Ohm's Law (V=IR) to give I = V/R = conductance(V), where V is voltage, I is the current (analogous to the leak), R is resistance and 1/R is conductance. The exponentially increasing non-Ohmic leakage (usually measured by O2 uptake in the presence of oligomycin) shown in the inserted graph in Figure 2, is not associated with non-specific membrane damage. A particular uncoupling membrane protein, UCP1, is especially prevalent in hibernators, small mammals, and human infants, which allows them to stay warm. The UCP family is activated by fatty acids, which may act as required cofactors for the protein or as antagonists of the inhibition of UCPs by nucleotide diphosphates. Two methods are used to determine the efficiency of coupling of oxidation and phosphorylation. • One is to determine the ADP/O ratio as both are substrates for oxphos, where O represents the amount of O2 consumed in the conversion of a defined amount of ADP to ATP when ADP is at high levels (which might not be physiologically relevant). This would give the maximum phosphorylation rate (defined as state 3). • A second is to measure ATP (or ADP)/O ratio directly through methods that give spectrophotometric signals such as the production of NADH by a coupled hexokinase/glucose-6-phosphate dehydrogenase assay in which ATP is used in the following reactions to make NADH, as shown in Figure \(3\). Figure \(3\): Generation of NADH on the combined reaction of hexokinase and glucose-6-phosphate dehydrogenase. In this coupled assay, one ATP used produces one NADH, which absorbs at 340 nm. Also, the ATP used as a substrate in the coupled reaction has left the mitochondria for it to have access to hexokinase, so the internal ratios of ADP/ATP in the respiring mitochondria are unchanged during the time course of the reaction. This method also allows the measurement to be made under nonsaturating ADP concentrations. Uncoupling of electron transport and ATP synthesis can occur through a combination of mechanisms, including proton loss at ANT and UCPs, at the complexes in electron transport that transport electrons and protons, and through ATP synthase itself. Other mechanisms would include substrate depletion or effects on the mitochondrial permeability transition pore (mPTP), which is an inner membrane transmembrane protein, which is gated open by increases in matrix Ca2+, adenine nucleotide decreases and increases in Pi. It is also a voltage-gated channel. When open the mitochondrial membrane depolarizes, ATP is depleted and cell death can be triggered. We'll discuss this complex later. Figure \(4\) shows the chemistry used to analyze the efficiency of oxphos and their graphical results. Figure \(4\): Measurement of coupling efficiency in respiring mitochondria. Eyenga et al. Ibid. Panel (A) shows the chemistry involved in the determination of the ATP/O ratio to measure coupling efficiency. Note that this step allows the regeneration of ADP and efficiency can be measured at any ADP concentration. Panel (B) shows changes in O2 with time. The respiration rate is given by the slope of the curve. Until substrate is added, O2 levels are constant. On the addition of substrate, O2 is consumed in electron transport so levels O2 levels fall linearly. When ADP is added, the rate of O2 consumption increases which is reflected in a steady linear decline in O2 levels. This state which is at fixed ADP levels (as illustrated in Panel (A)), is called state 3 (maximum ATP synthesis at a given ADP level). When oligomycin, an inhibitor of proton flow across ATP synthase which inhibits ATP synthesis (and ADP use) is added, state 4 ensues with shows decreased O2 consumption when ATP synthesis is stopped. Note the parallel lines for stages 2 and 4. The Cytochrome c oxidase activity at the end of the curve is the maximal respiration with ascorbate (as an oxidizing agent) and N,N,N′N′-tetra methyl-p-phenylenediamine (TMPD) as a substrate. Panel (C) shows a graph of the rate of ATP synthesis vs the rate of O2 consumption. The slope gives the rate of ATP generation to that of O2 consumption (ie. the ATP/O ratio). Note that the graph is linear:O is linear over a range of fixed, non-saturating ADP concentrations. Changes in Proton Gradient from "Slippage" Slippage in the electron transport chains: Redox Slipping Proton translocation from the matrix to the inner membrane space is an integral part of the function and activity of the electron transport chains that establish the pH gradient and hence the Δp. These components are Complexes I, III, and IV. Complex IV (cytochrome C oxidase - CCO) appears to be especially important. Protons are removed by CCO from the matrix on the reduction of O2 to H2O. This requires exactly 4 electrons and 4 protons so the stoichiometry is fixed. Also, CCO moves protons into the inner membrane space in the formation of the pH gradient. This is where the electron-to-proton ratio may vary depending on redox and coupling conditions. Alterations in CCO or even decreases in its concentration would impact the movement of protons as well. Instead of calling these alterations in nominal proton translocation leakages, they are called slippage since an ordinary function of CCO is proton translocation. Slippages through CCO would decrease proton transport and decrease ATP/O. Reactive nitrogen species (NO and peroxynitrite) can also decrease ATP/O. Slippage at ATP synthase Proton leakage at ATP synthase would also result in a decreased ATP/O ratio. This occurs especially if nucleotides are low or if the holoenzyme is compromised by low subunit availability. ATP synthase forms dimers and other multimers to form supercomplexes, and this affects (or is determined by) the formation of invaginations in the inner membrane which form cristae. Given that the multimeric c-ring of F0 effectively moves protons across the inner membrane, it is a likely source of slippage. It most likely occurs through a modified version of the F0 ATP synthase unit called the ATP synthase c-subunit leak channel (ACLC). This is different from the mitochondrial permeability transition pore (mPTP) mentioned above. Alternatively, it may be that the ACLC might be part of the leak channel of the mitochondrial permeability transition (MPT) pore discussed above. Studies show that the c-ring can form large conductance channels that are voltage-gated without regulatory subunits. In addition, added F1 subunit inhibits it. In neurons stimulated by the excitatory neurotransmitter glutamate, F1 dissociates from the F1F0 complex, allowing the c-ring to slip protons through the membrane. Finally knockdown of the c-subunit eliminates high conductance. Hence the c-ring is part of a type of mPT that is regulated by cyclophilin D (CypD), which is a mitochondrial peptidyl-prolyl cis-trans isomerase, that regulates the mitochondrial permeability transition pore (PTP). In vitro, c-rings incorporated into vesicles form a large voltage-gated channel that is inactivated by the addition of F1. Also in neurons stimulated with the excitatory neurotransmitter glutamate, which leads to increases in cytoplasmic Ca2+, binding and ensuring conformational changes in the complex leads to dissociation of F1 from the complex, freeing F0 for proton "slippage" and transport uncoupled from ATP synthesis. A Proposed “bent-pull-twist” model of ACLC gating in physiological and pathological conditions is shown in Figure \(5\). Figure \(5\): Bent-pull-twist model of ACLC gating in physiological and pathological conditions. Mnatsakanyan, N., Park, HA., Wu, J. et al. Mitochondrial ATP synthase c-subunit leak channel triggers cell death upon loss of its F1 subcomplex. Cell Death Differ (2022). https://doi.org/10.1038/s41418-022-00972-7. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/. Panel A shows c-ring diameter changes during the electrophysiology recordings, upon the application of voltage and F1. F1 added during c-ring recordings probably inactivates the channel due to the specific interactions between F1 and c-ring that induce twisting motions in c-subunit monomers in a clockwise direction, to stabilize the ring, reduce the pore diameter and close the channel. Panel B shows reversible brief openings of ACLC in physiological conditions. Panel C shows that non-reversible dissociation of F1 from FO occurs during long-lasting openings of the c-ring channel in severe pathology. For simplicity, only ATP synthase monomer is shown. ATP synthase subunits are drawn as ribbon representations (modified PDB ID code: 6J5I) [14]. In B and C, red arrows indicate the path of ion flow through the channel. Closed and open conformations of the channel are noted. A closer look at uncoupler protein 1 (UCP1) UCP1 is an inner membrane protein in the mitochondria that bypasses the movement of protons through the c-ring of F1. UCP1 is activated by fatty acids and inhibited by purine nucleotides. The mechanism of proton translocation is unclear. It could arise from a "handshake" of protons through a series of linked hydrogen bond donors and acceptors in the membrane protons. Evidence suggests that what moves are bound and protonated fatty acids which flip across the inner membrane where the carboxyl group deprotonates in the high pH matrix side of the membrane. The role of UCP1 in this process could be to flip the deprotonated fatty acid back across the membrane. This model is called the protonophoretic model in which the fatty acid acts as a "protonophore" much like valinomycin, which transports K+ across the membrane and is called an ionophore. UCP1 doesn't interact with the H+ directly but just flips the deprotonated fatty acid back to continue the cycle. This model would give a stoichiometry of one flipped negatively charged fatty acid per proton transferred. In the second "H+-shuttling" model, UCP1 is considered a fatty acid/H+ symporter (carries both species in the same direction). In this model, a fatty acid is strongly bound to UCP1 with the head group inside a cavity in the protein. Both models required UCP1 to carry deprotonated fatty acids back across the membrane. The NMR structure of a similar protein, UCP2, is known. It can bind fatty acids with the acyl tail near the matrix side. However, UCP2 does not seem to be involved in thermogenesis and doesn't uncouple oxphos. It binds and catalyzes the exchange of malate, oxaloacetate, and aspartate for phosphate and H and hence has other metabolic roles. UCP 1 and 2 are about 60% homologous but have similar alpha-helical structures based on modeling. Perturbation of NMR signals from assigned side chains was used to show the fatty acids bind to UCP1 and it's closer to the matrix than the inner membrane space. Two key lysines (K56 on helix 1 and K269 at the matrix side of helix 6) probably bind with the carboxylate end of the fatty acid. UCP1also has a hydrophobic grove (T31, F32, L34, D35, L278, G279, S280, W281, and V283) to accommodate the fatty acid. In addition, both UCP1 and UCP2 transport/flip alkyl sulfonates, analogs of fatty acids. This is consistent and required with both models. Figure \(6\) shows an interactive iCn3D model of the predicted AlphaFold model of human UCP1 (P25874). The subunits are shown in the same color as Figure 1 above. Figure \(6\): Predicted AlphaFold model of human UCP1 (P25874). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...Qf8W1MgVH6u546 The blue colors represent assigned structures with the highest predicted confidence, and the red the lowest. The amino acid side chains that are implicated in binding to fatty acids based on perturbation of NMR signals on fatty acid binding (http://dx.doi.org/10.1016/j.str.2017.07.005) are shown in sticks, with CPK colors and labeled. The membrane would be positioned perpendicular to the average vertical orientation of the helices in the above orientation. Additional Controls From the perspective of moving electrons into the electron transport pathway, the key deliverers of electrons are NADH derived from integrated multiple pathways at Complex I (as illustrated in Figure 1) and FADH2 from the citric acid cycle at Complex II. Hence if intermediates in the citric acid cycle are low, the flux through electron transport is compromised. Flux through Complex I seems to determine the rate-limiting steps for oxygen use. In the previous chapter section, we calculated the total chemical potential driving force across the membrane. A pH difference of 1.4 across the membrane provides a -1.9 kcal/mol (-8 kJ/mol) H+ chemical potential compared to the -3.23 kcal/mol (13.5 kJ/mol) from the transmembrane potential, Δψm. Both contribute to the thermodynamically favorable movement of H+ across the inner membrane in respiring mitochondria, but Δψm is a large contributor. Hence Δψm and ATP synthesis are also coupled with maximal synthesis occurring between -100 to -150 mV. At values around -200 mV, give rise to increased permeability for protons (which decreases Δp) but also the production of superoxide and peroxides. Hence Δψm is also regulated. ADP levels control ATP synthesis as well, with higher concentrations leading to increased ATP synthesis and a decrease in Δp. Decreased ATP synthesis from increased Δp affects coupling, slippage in cytochrome C oxidase, and heat generation. Reduction of O2 to water by stepwise addition of electrons involves the generation of superoxide, O2-. As shown in an earlier section, superoxide formed by Complex I would move to the matrix, while those formed by Complex III to both sides of the membrane. Hydrogen peroxide can also form from monoamine oxidase and α-ketoglutarate dehydrogenase, which produces increased peroxides with increasing NADH concentrations. Respiratory state 4 (when proton transport is blocked by oligomycin) leads to higher levels of ROS. Hence ROS levels are also determined by not only the respiratory state but also Δp. Other control mechanisms are potentially derived from other protein subunits that bind to the complex within a lipid bilayer. These include A6L, DAPIT (diabetes-associated protein in insulin-sensitive tissues), and the proteolipid 6.8PL. Another is the ATPase inhibitory factor 1 (IF1), which turns out to be a significant regulator. When the Δψm falls too low, IF1 inhibits ATP depletion which leads to ATP hydrolysis by the complex to move protons from the matrix to the inner membrane space. The interaction of IF1 with ATP synthase is shown in Figure \(7\). Figure \(7\): Structure of the bovine H+-ATP synthase and binding site of IF1. Esparza-Moltó, P.B., Nuevo-Tapioles, C. & Cuezva, J.M. Regulation of the H+-ATP synthase by IF1: a role in mitohormesis. Cell. Mol. Life Sci. 74, 2151–2166 (2017). https://doi.org/10.1007/s00018-017-2462-8. Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/) Panel (a) shows the soluble F1-ATPase domain composed by the 3α3β subassembly (salmon/red) and γ (dark blue), δ and ε (light blue) subunits, while the membrane-embedded Fo domain is formed by subunit a (red) and a ring of 8c subunits (light/dark blue). Both domains are linked together by a central stalk (γ, δ, ε subunits of the F1 domain) and a peripheral stalk (b, d F6, A6L, OSCP subunits, in orange). The 3D structure of the peripheral stalk is not fully resolved. Except for a and A6L subunits, the remainder is encoded in the nucleus. (model comprised of multiple PDB structures) Panel (b) shows the lateral and basal view of the bovine F1 domain (α subunit is shown in salmon, β in red, and γ in blue) complexed with a fragment of IF1 (green). IF1 binds to the αβ interface through residues 1–37 and also contacts the γ subunit. (PDB: 1OHH) IF1 has an intrinsically disordered N-terminal domain that binds to ATP synthase between the α and β subunits of the F1, where it adopts an alpha-helical conformation. It can dimerize through self-interactions with the C-terminal domain. Interestingly, bound IF1 inhibits both ATP synthesis and hydrolysis as well as H+ translocation through the complex. These activities are illustrated in Figure \(8\). Figure \(8\): IF1 inhibits H+ translocation through the H+-ATP synthase in both synthetic and hydrolytic modes. Esparza-Moltó, ibid. Panel (a) shows H+ uptake is induced by valinomycin-mediated K+ release from FoF1-K+ liposomes with the H+-ATP synthase functioning in the hydrolytic mode. Panel (b) shows H+ release is induced by valinomycin-mediated K+ uptake in FoF1-K+ liposomes with the H+-ATP synthase functioning in the synthetic mode. The rates of H+ uptake (a) and H+ release (b) are reduced when the liposomes are incubated with increasing concentrations of isolated IF1. Black circle valinomycin. Five key histidines in IF1 play a role in the inhibition of the ATPase activity and the oligomerization of IF1. When the pH gradient collapses, and the matrix becomes more acidic (as seen in hypoxia and ischemia), IF1 form dimers that inhibit ATP hydrolysis. At higher pHs, IF1 forms a dimer of IF1 dimers, or effectively a tetramer, which prevents its binding to ATP synthase and hence prevents the inhibition of ATP synthesis. An IF1-H49K mutant is an active inhibitor even at higher pHs. These properties are illustrated in Figure \(9\). Figure \(9\): Post-translational regulation of IF1 activity. Esparza-Moltó, ibid. IF1 inhibition of ATP hydrolysis is not only relieved by pH-dependent oligomerization as illustrated above, but also by phosphorylation by a mitochondrial Protein kinase A-like molecule, or by its interaction with yet another protein, BP. A conserved serine 39 is important in its inhibitory action. When dephosphorylated, IF1 binds to ATP synthase and inhibits both ATP synthesis and hydrolysis. When phosphorylated it can't bind and hence can't inhibit ATP synthase. Mitochondria - Nuclear Signaling We would be remiss to not include the role of the mitochondrial genome and its relationship to the nuclear genome in the control of mitochondrial function, including oxidative phosphorylation. Mitochondrial arose from the internalization of a distant bacteria into a eukaryotic cell. They developed an endosymbiotic relationship in which some of the genes from the original bacterial species remain in mitochondria today. The mitochondrial genome is shown in Figure \(10\). Figure \(10\): Mitochondrial genome. https://commons.wikimedia.org/wiki/C...ial_DNA_en.svg Mitochondrial DNA is a double-stranded molecule of 16.5 kilobases. It encodes some proteins, as well as its own tRNA and ribosomal RNAs. There are many copies (100s to 1000s) of the circular genome in each cell. Some (actually 13), but not all of the genes for proteins required for oxidative phosphorylation are found in the mitochondrial genome. The rest reside in the nuclear genome, and after transcription and translation, the resulting protein must be imported into mitochondria. In fact, over 95% of mitochondrial proteins are encoded by nuclear genes. Mitochondria move in the cells guided by the internal cytoskeleton to sites where energy (ATP) is needed. They also participate in buffering intracellular Ca2+ ions in the cell, much like the ER with which mitochondria form intimate membrane contacts. The ER also makes contact with the outer nuclear membrane, so, likely, the mitochondria can closely associate with the nucleus as well. Hence there must be regulation of the respective genomes to produce the correct type and amounts of proteins required for oxidative phosphorylation. To accomplish this, there is a bidirectional regulated signaling between the two organelles. There is direct signaling between the nucleus and mitochondria and retrograde signaling in the other direction. Signals and processes affected in these signaling pathways are shown in Figure \(10\). jdjkfj Figure \(11\): Signaling from the mitochondria to the nucleus. Brittni R. Walker and Carlos T. Moraes. Biomolecules 2022, 12(3), 427; https://doi.org/10.3390/biom12030427. Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/) Signals originating from the mitochondria, including calcium, reactive oxygen species (ROS), and AMP/ATP, often stimulate pathways, leading to transcriptional changes in the nucleus. Nuclear responses can involve upregulating cell proliferation and anti-apoptotic factors, as well as proteins involved in mitogenesis, such as PGC-1α and nuclear-encoded mitochondrial proteins. The levels of other molecules, such as TCA intermediates Acetyl CoA and α-ketoglutarate, can influence the epigenome by modifying methylation and acetylation, and consequently, the cell physiology. We've encountered several key enzymes involved in the regulation of metabolism, including: • AMP-activated protein kinase (AMPK) senses the energy state (AMP/ATP ratio) of the cell. When ATP is low (AMP is high), it activates pathways to produce ATP and inhibits pathways that require ATP. AMPK has been found at or near the mitochondrial membrane and may also be found in it. AMPK also promotes mitochondrial biogenesis (mitogenesis) • mTOR (mammalian target of rapamycin ) which is a serine/threonine kinase that regulates protein synthesis and other anabolic pathways based on the energy status of the cell. processes in response to growth factors, energy status, and oxygen levels. It also simulated mitogenesis.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/19%3A_Oxidative_Phosphorylation/19.03%3A_Regulation_of_Oxidative_Phosphorylation.txt

Search Fundamentals of Biochemistry Introduction We have seen how we can transduce the chemical potential energy stored in carbohydrates, into the chemical potential energy of ATP. This occurs namely through coupling the energy released during the thermodynamically favored oxidation of carbon molecules through intermediaries (high energy mixed anhydride in glycolysis or a proton gradient in aerobic metabolism) to the thermodynamically uphill synthesis of ATP. There is a situation that occurs when we wish to reverse the entire process: $\ce{CO2 + H2O → carbohydrate + O2} \nonumber$ This process is, of course, photosynthesis, which occurs in plants and certain photosynthetic bacteria and algae. Given that this process must by nature be an uphill thermodynamic battle, let us consider the major requirements that must be in place for it to occur: • A strong oxidizing agent must be formed which can oxidize water to dioxygen. We know that redox reactions occur in the direction of a stronger to a weaker oxidizing agent (just as acid/base reactions are thermodynamically favored in the direction of strong to weak acid). Somehow we must generate a stronger oxidizing agent than product dioxygen, which often has the most positive standard reduction potential in common tables. • Plants must have high concentrations of a reducing agent for the reductive biosynthesis of glucose from CO2. The reducing agent used for most biosynthetic reactions in nature is NADPH, which differs from NADH only by the addition of a phosphate to the ribose ring. This phosphate differentiates the pool of nucleotides in the cells used for reductive biosynthesis (NADPH/NADP+) from those used for oxidative catabolism (NADH/NAD+) • Finally, plants need an abundant source of ATP which will be required for reductive biosynthesis. The light reaction of photosynthesis produces these three molecules, O2, NADPH, and ATP. We will explore the light reaction in the next few sections of this chapter. The dark reaction, which as the name implies can occur in the dark, involves the actual fixation of carbon dioxide into carbohydrates using the ATP and NADPH produced in the light reaction. The energy to power the light reactions comes directly from sunlight. Clue two is that plants have an organelle that animal cells don't - the chloroplast. Its structure is in many ways similar to mitochondria in that it has an outer membrane, an intermembrane space, and an inner membrane. In addition, it has a series of stacked, interconnected compartments called thylakoids bounded by a thylakoid membrane surrounding a lumen. A schematic is shown in Figure $1$ below. Stacks of thylakoids are called grana. Each thylakoid represents one granum. The thylakoids are where the light reactions occur. In contrast, the stroma contains the enzymes for the dark reactions of photosynthesis. The enzyme complexes involved in the light reaction are aligned in the thylakoid membrane just as the membrane complexs involved in mitochondrial electron transport/oxidative phosphorylation were aligned in the mitochondrial inner membrane. That process used Complex I, Complex III, Complex II (succinate dehydrogenase), and Complex IV to transfer electrons from NADH and FADH2 to increasingly potent oxidizing agents (ubiquinone, cytochrome C) ending with dioxygen. The energy released during that thermodynamically favored process was captured in the formation of a proton gradient, which collapsed through the F0F1ATPase to drive ATP synthesis. For the light reaction, three complexes, the Light Harvesting ComplexII -Photosystem II (also called the LHC-PSII supercomplex), cytochrome b6f, and the light-harvesting complex Photosystem I (called the LHC-PSII supercomplex) are used to carry out the light reactions of photosynthesis. These include the photooxidation of water to produce O2 and the transfer of the lost electrons through a series of mobile electron carriers (plastoquinone and plastocyanin) to a terminal acceptor, NADP+ to form the reducing agent NADPH needed for carbohydrate synthesis in the dark reaction. The energy released during the electron transfer process is likewise captured in the form of a proton gradient as protons are moved from the chloroplast stroma to the lumen of the thylakoid. The collapse of the resulting proton gradient powers ATP synthesis. These reactions are shown in Figure $2$. In this section, we will explore the absorption of light by the light-harvesting complex (LHCII) of the LHC-PSII supercomplex. Before we get into too much detail, let's start with a simplified review of light absorption in the LHCII. Absorption of Light Plants have many pigments (chlorophylls, phycoerthryins, carotenoids, etc.) whose absorption spectra overlap that of the solar spectra. The main pigment, chlorophyll, has a protoporphyrin IX ring (same as in heme groups) with Mg2+ at its center instead of Fe2+. When the chlorophyll absorbs light, the excited electrons must eventually relax to their ground state. It can do this by either radiative or nonradiative processes. In radiative decay, a photon of lower energy is emitted (after some energy has already been lost by vibrational transitions) in a process of either fluorescence or phosphorescence. In nonradiative decay, the energy of an excited electron can be transferred to another similar molecule (in this case other chlorophyll molecules) in a process that excites the electron in the second molecule to the same excited state. It is as if a photon is released by the first excited molecule, which then is absorbed by an electron in a second molecule to excite it to the same excited state. However, no photon is involved in the energy transfer. In this fashion, energy is transferred from one chlorophyll to another. This type of energy transfer is called resonance energy transfer or exciton transfer, as shown in Figure $3$. Because of its unique environment, one type of chlorophyll has slightly different characteristics. The energy level of the first excited state in the chlorophyll reaction center is lower than in the rest of the chlorophyll molecules, in much the same way that pKa values of amino acid side chains differ with the local and solvent environment, and the standard reduction potential of FAD molecules that are tightly bound to enzymes differ due to the different environment of bound FAD/FADH2. Instead of a radiationless transfer of energy to this special chlorophyll, an actual electron from the excited state chlorophyll is transferred, which by definition is a redox reaction. The electron donor (the excited chlorophyll) loses an electron (an oxidation reaction) as the recipient molecule gains one (a reduction). This charge (electron) transfer reaction produces charge separation in the formation of a positively charged chlorophyll (now an oxidizing agent) and a negatively charged chlorophyll (now a reducing agent). The chlorophylls directly involved in this final process are collectively called the reaction center. This process is shown in Figure $4$ below. The reaction center chlorophylls absorb light at 680 nm so sometimes these chlorophylls are labeled P680. There are 4 unique chlorophylls (PD1, PD2, ChlD1, and ChlD) that are the main players in the reaction center. Both sets of labels are shown in the figure above. Figure $5$ shows a cartoon of the absorption of photons and subsequent handoff of energy to "antennae" chlorophylls leading to photoexcitation of the reaction center and subsequent transfer of an electron to the special chlorophyll which has a lower first excited state energy. This molecule is named pheophytin A. It is identical to chlorophyll A but lacks the central Mg2+ ion. Photosystems I and II contain many chlorophyll molecules that act as antennas that transfer energy to the reaction centers. The "antenna" proteins involved in photon adsorption and energy transfer in Photosystems I and II are shown in Figure $6$ below. Note the D1, D2, cp47, and cp43 protein subunits in the figure below are also shown in the figure above. The D1 and D2 subunits contain the reaction center chlorophylls. The different chlorophylls have absorption spectra that overlap reasonably well with the solar spectrum, as illustrated in Figure $7$. The black lines show the solar flux spectrum (photons per meter square per wavelength) from 300 to 1200 nm. The yellow line is the photon energy (E=hc/λ) at each wavelength. The absorbance spectra of the chlorophyll species are normalized to the same maximal value. If you use the energy required to make glucose from CO2 and H2O as a standard, about 95% of the incoming solar energy is wasted since not all of the incident light photons have the right wavelength. Waste also occurs due to reflectance and heat generation. The Light Harvesting Complex (LHCII) - Photosystem II (PS II) Supercomplex Now let's look in more detail at the chloroplast thylakoid membrane complex that interacts with light and results in the oxidation of water to form O2. This first structure is called the Light Harvesting Complex II (LHCII) - Photosystem II (PS II) Supercomplex. It is a super complex (a pun) to understand. The supercomplex has a PSII core complex interacting with a variable number of light-harvesting complex IIs (LHCIIs) complexes. Sometimes the entire super complex is more simply called Photosystem II. The supercomplex consists of • a PSII core C with a Mn4CaO5 cluster called the oxygen-evolving complex (OEC or OEX) that oxidizes water to O2; • peripheral antennae complexes M and S, also called light-harvesting complexes II (LCHIIs) The core part of the Photosystem II supercomplex contains the key to the existence of aerobic organisms as it produces the dioxygen in our atmosphere. Its chemistry is phenomenal. Many are seeking to modify it to produce, in addition to O2, a green energy source, H2. We will explore the mechanisms for the oxidation of water to O2 by a PSII-bound inorganic Mn4CaO5 cluster, the oxygen-evolving complex (OEC) in the next section. Let's first look at PSII in light of the discussion above. Where are all the chlorophylls? The mystical "reaction center"? The PSII Core Complex (within the supercomplex) has over 20 subunits. These include: • reaction center subunits D1 and D2 • inner antennae subunits CP43, CP47 • many other small protein subunits • peripheral subunits that project into the lumen that interact with the OEC The peripheral antennae complexes (light-harvesting complexes - LHCIIs M and S - consist of proteins encoded by 6 different Lhcb genes (1-6). • Lhcb1-3 monomers form trimeric (homo- or hetero) LHCIIs (Lhcb1-3). An example is (Lhcb3)3. That trimer is often called the M-LHCII. There is also an S-form trimer as well as L- and N-forms. S represents a strong association with the reaction center, M for moderate and L for loose. • Lhcb4-6 are called the minor components and consist of the monomeric proteins named CP29, CP26, and CP24, respectively. Light-harvesting complex proteins Each Lhc apoprotein consists of 3 alpha-helices and binds 8 chlorophylls a, 6 chlorophyll b, and 4 carotenoid molecules. Figure $8$ shows an interactive iCn3D model of the monomeric pea chlorophyll a-b binding protein AB80 (LHCII type I CAB-AB80) (2BHW) Figure $8$: Monomeric pea chlorophyll a-b binding protein AB80 (LHCII type I CAB-AB80) (2BHW). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...QtEEWeQ5Y9n2i8 The color scheme is as follows: • protein - secondary structure colors • Chlorophyll a - Green • Chlorophyll b - yellow • carotenoids - cyan Now let's look at a trimeric form of the same protein. of LHCII, the chlorophyll-binding protein, from PSII. It's also called LHCII type I CAB-AB80. We showed the role of the LHCII trimer in the figures above. The subunits function to absorb light for the light reaction and through resonance transfer energy to the reaction centers in PSII and in addition PSI. Figure $9$ shows an interactive iCn3D model of the trimeric Light Harvesting Complex (LHC_II) from pea photosystem II (2bhw). Figure $9$: Light Harvesting Complex (LHC_II) from pea photosystem II (2bhw). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...BK6WvzrxABPDh6 The protein is shown in grey. To simplify the model, chlorophyll A molecules are shown in magenta, and chlorophyll B isomeric variants are in green. Note a large number of chlorophylls (42) in the LHCII complex, which differentiates it from most protein complexes which might have just a few ligands bound. Photosynthetic bacteria and plants have an abundance of molecules that interact with visible light. The main one found in PS II is chlorophyll a whose structure is shown in Figure $10\ below, along with the variant, chlorophyll b. Remember the pheophytin A is identical to chlorophyll A but lacks the central Mg2+ ion. Photosystem II (PS II) Supercomplex Structures The most common version of the supercomplex is a dimer. A simplified cartoon of the monomeric version of the PSII-LHCII supercomplex is shown in Figure \(11$. The PSII core is shown in the blue rectangle. The CP29, CP26, and CP24 in the peripheral antennae complexes (light-harvesting complexes - LCHIIs) are shown outside of the blue-outlined rectangle. They surround the PSII core. • The PSII core C (outlined in the blue box) contains two 2 reaction center subunits D1 and D2, as well as two inner antennae subunits CP43 and CP47. The core also contains extrinsic subunits (pink) surrounding the OEC (red spacefill) • A Strongly bound LHCIIs protein, S -LHCII, left (maroon rectangle) interacting with CP26. The S-LHCii complex is a trimer of individual Lhcb monomers. • A Moderately bound LHCIIs protein complex, M-LHCII, right (lighter blue rectangle) interacting with the CP24/CP29 dimer. The M-LHCII complex is a trimer of 3 lcbh-3 subunits (Lcbh3)3. There can be a variable number of LHCIIs in the complex. The predominant form of the supercomplex is the C2S2M2 complex. There are also many other proteins in the supercomplex that are not shown in Figure 11. A cartoon showing the top-down view of the actual supercomplex dimer, C2S2M2, is shown in Figure $12$. You could imagine forming the dimer by spinning the monomer CSM 1800 and reproducing the monomer there. Figure $13$ shows an interactive iCn3D model of the full C2S2M2-type PSII-LHCII supercomplex from Pisum sativum (5XNL). (long load time) Figure $13$: C2S2M2-type PSII-LHCII supercomplex from Pisum sativum (5XNL). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...F4WFfHoevxaVV8 (long load time) Only a single CSM monomer of the C2S2M2 dimeric supercomplex is colored. The colors match those in Figures 8 and 9. The molecules shown in gray sticks that outline a bilayer include chlorophyll Bs (98 of them), chlorophyll As (216), and carotene or its derivatives (88) as well. The different proteins in the supper complex are colored as shown below. • CP47 Rx Center (inner antenna protein) Dark Blue • CP43 Rx Center (inner antenna protein) - Purple • D1: green • D2: yellow • OEC: spacefill Red • O, P, Q - OEC peripheral: Light pink • CP24 (peripheral antenna protein): yellow • CP29 (peripheral antenna protein): orange • CP26: Light blue • LHCIIs (S and M) - not shown • membrane bilayers - cyan spheres (A quick load version of the C2S2M2-type PSII-LHCII supercomplex with different coloring than above) PSII Supercomplex Regulation Plants have evolved a great ability to absorb light over the entire visible range of the spectra. Can they absorb too much energy? The answer is yes, so plants have developed many ways to protect themselves. IF too much light is absorbed, the pH gradient developed across the thylakoid membranes becomes greater. This is sensed by a protein, PsbS, and through subsequent conformational changes transmitted through the light-harvesting antennae, the excess light energy is dissipated as thermal energy. Mutants lacking PsbS showed decreased seed yield, a sign that it became less adaptable under conditions of stress (such as exposure to rapidly fluctuating light levels). Molecules called xanthophylls and other carotenoids such as zeaxanthin are also important in excess energy dissipation. These molecules appear to cause excited state chlorophyll (a singlet-like excited state dioxygen) to become deexcited. Without the xanthophylls, the excited state chlorophyll could deexcite by transfer of energy to ground state triplet dioxygen, promoting it to the singlet, reactive state, which through electron acquisition, could also be converted to superoxide. These reactive oxygen species (ROS) can lead to oxidative damage to proteins, lipids, and nucleic acids, alteration in gene transcription, and even programmed cell death. Carotenoids can also act as ROS scavengers. Hence both heat dissipation and inhibition of the formation of ROS (by such molecules as vitamin E) are both mechanisms of defense against excessive solar energy Given that both plants and animals must be protected from ROS, antioxidant molecules made by plants may prove to protect humans from diseases such as cancer, cardiovascular disease, and general inflammatory diseases, all of which have been shown to involve oxidative damage to biological molecules. Humans, who can't synthesize the variety and amounts of antioxidants that are found in plants, are healthier when they consume large amounts of plant products. These phytomolecules also have other properties, including regulation of gene transcription which can also have a significant effect on disease propensity. Plants have to respond to different qualities and quantities of light. When light is low, they seek to maximize light capture. Too much light could damage a plant so molecular adaptions are made to prevent it. Part of the regulation occurs in the stoichiometry of the supercomplex by altering the antenna protein (the LHCs) composition. The most abundant component of the complex is C2S2M2 and C2S2M. C2S and C2S2 increase as an adaption to increasing levels of light. Light levels can also promote grana membrane association mediated by interactions of a PSII-LHCII supercomplex (PSII-LHCIIsc) on one thylakoid membrane interacting with a PSII-LHCIIsc on an adjacent thylakoid membrane to form a large PSII-LHCIIsc dimer. The structure and properties of paired PSII-LHCIIsc are illustrated in Figure $14$. The figure also shows the variation in the PSII–LHCIIsc with light intensity. At low light C2M2S2 prevails while at high light intensity C2S2 is most abundant. The system moves to regulate activity by increasing the abundance of LHCIIs in low light and decreasing them in high light intensities! Figure 1 shows the stacking of individual granum in the chloroplasts. The stacking is maintained at various light levels and is mediated by loops of the LHCII trimers that are exposed in the stroma and Lhcb4 subunits on adjacent membranes. The stromal surfaces are flat and tightly stacked in grana. Stacking is a dynamic process and depends on cross-membrane interactions between and reorganization of the PSII -LHCIIsc. The PSII–LHCIIsc is regulated through light-dependent post-translational phosphorylation, particularly on LHCII, and acetylation. which further regulates energy distribution. In plants, the core proteins CP43, D1 and D2, and PsbH are phosphorylated by the kinase Stn8. The peripheral antenna LHCII proteins are phosphorylated by Stn7. Phosphorylation of PSII is not seen in cyanobacteria and red algae. Higher light levels promote higher levels of core protein phosphorylation. Stn7 appears to be inhibited at high light levels. Figure $15$ shows putative Stn8 phosphorylation sites in the PSII -LHCIIsc The structure is a model pieced together from multiple pdb files. The approximate positions of phosphorylation sites of D1, D2, CP43, PsbH, and CP29 are shown. Most free N-terminal loops in LHCII can be phosphorylated. Both phosphorylation and lysine acetylation on Lhcb2 N-terminal loops help regulate the redistribution of LHCII from PSII (in grana stacks) to PSI (in single-layered thylakoid regions). Grana stacking by N-terminal loop association between facing PSII–LHCIIsc and their N-terminal acetylation appear to strengthen grana stacking. How does PSII respond when not the intensity but the wavelength of light is changed? For this, we have to briefly discuss photosystem I (PSI ), which we will explore in more detail in Chapter 20.3. Both PSII and PSI work together to transduce light into chemical energy so you would expect that their activities are regulated in a linked fashion. They have different absorbance spectra characteristics as well, with PSI absorbing more in the red region. If the effective absorbance (normalized for concentrations and LHCIIs) were the same, you would predict that the effective absorbance ratio over a broad wavelength range for the two photosystems, PSI/(PSI+PSII) would be 0.5. This is approximately the case over the entire spectral wavelength except for between 670-730 nm, where the ratio is close to 1, showing that PSI absorbance and hence activity is tilted toward the red end of the absorbance spectra. Changes in light characteristics (i.e. wavelength) would then affect each photosystem differently and can cause imbalances in their activities and their states, which should lead to a restorative balance. When exposed to far-red light, the systems move to state I. In this state, the major mobile antenna proteins (LHCIIs) move to PSII to restore a "photoabsorption" balance. When exposed to light depleted in the high end of visual spectra, the system moves to state II, in which mobile LHCIIs move to PSI. What an interesting reciprocal way to balance activity, even if it is hard to conceptualize regulation involving the movement of membrane proteins. Of course, such movement is seen often in the clustering of ligand-bound membrane receptors. As mentioned above, the location and hence movement of the LHCII is regulated by phosphorylation by LHCII kinase. We'll explore that more in Chapter 20.3 when we discuss PSI in more detail. A closer view of the reaction center and its local environment Let's look at the structure of a simpler PSII complex from Thermostichus vulcanus (3WU2). It has 70 chlorophyll a molecules, 4 special chlorophylls, 4 pheophytin As (PHOs), 20 beta-carotenes, 4 plastoquinol-9s (PL9s), 4 hemes, and 2 caroten-3-ols. Figure $16$ shows the arrangement of chlorophyll molecules in the CP47 and CP44 (not CP43 as shown in several of the above diagrams) antennae subunits of PSII from T. vulcanus (3WU2). . • CP44 is the Photosystem II CP44 reaction center protein, psbC gene, Photosystem II 44 kDa reaction center protein; • CP47 is the Photosystem II CP47 reaction center protein, psbB gene; Photosystem II CP47 chlorophyll apoprotein Sandwiched in between them is the reaction center containing the 4 special chlorophylls, the inorganic metal cluster called the oxygen-evolving complex (OEC or OEX), and a key amino acid near the OEC, Tyr 161 (pdb 3ARC). Two special chlorophylls, ChlD1 and ChlD2, accompanied by partner chlorophylls, PD1 and PD2, that are coplanar to each, are found near the OEC and are the key chlorophylls in the reaction center that turn OEC into a powerful enough oxidant to oxidize H2O A special reaction center chlorophyll/pheophytin absorbs a photon of light at 680 nm so the species that absorbs the photon is given the label P680. On absorption, it forms the excited state, P680*. This transfers the excited state electron to pheophytin which forms a pheophytin radical anion, while the electron donor P680* becomes P680.+, a radical cation. The radical cation, an unstable species, can oxidize another molecule to regain stability, and in a series of addition linked oxidation stems, water is oxidized (O in water has an oxidation state of 2-) to O2 (in which each oxygen has an oxidation state of 0). This happens through the oxygen-evolving complex, which we will explore in the next section. Which of the chlorophyll molecules absorbs the light (ie. which is P680)? The answer is probably all four chlorophylls in the reaction center, PD1, PD2, ChlD1, and ChlD2 through a delocalization of the excited electron, which would be described by a wave function for the combined chlorophylls. On donation of the electron from the P680*, the positive charge, which can be also described as a "hole" (similar to transistors) delocalizes as well. Quantum mechanical calculations show a coupling between PD1 and PD2, such that 80% of the positive charge and radical character is situated on PD1 and 20% is on PD2.) Figure $17$ shows the four chlorophylls in the reaction center of T. Vulcanus. PD1 appears closest to the OC. Figure $18$ shows an interactive iCn3D model of the key chlorophylls and OEC of photosystem II from Thermostichus vulcanus (3WU2) Figure $18$: Key chlorophylls and OEC of photosystem II from Thermostichus vulcanus (3WU2). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...qYxWN5sKcquGK7 Now in any enzymatically catalyzed reaction mechanism, the enzyme must return to the beginning state and a path for electron flow must be apparent. How does the radical cation P680.+ return to the ground state? As we will see in the next section, it "grabs" an electron from the nearby Y161 (also called YZ), which then forms a radical cation, Y.+. This likewise returns to the ground state by grabbing an electron from the OEC which ultimately grabs one from water. This process repeats four times to remove the four electrons from two waters needed to form dioxygen. Figure $19$ shows the first step in process of reforming P680 and the formation of the Y161 radical cation. A different process occurs to remove electrons from the radical anion, P680.-. This is passed on to a series of other electron acceptors/carriers as part of the Z scheme for the light reaction of photosynthesis. The electron is ultimately passed on to NADP+ to form NADPH for the reductive biosynthesis of carbohydrates. Light is also absorbed in photosystem I, which does not form O2. Rather it receives electrons from a mobile electron carrier and passes them on NADP+ to form NADPH, a reducing agent needed for the reductive biosynthesis of carbohydrates. It also helps produce a proton gradient which helps drive ATP synthesis. With this background, we can now explore in greater detail the key reactions that enable the evolution of aerobic organisms.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/20%3A_Photosynthesis_and_Carbohydrate_Synthesis_in_Plants/20.01%3A_Light_Absorption_in_Photosynthesis_-_An_Overvie.txt

Search Fundamentals of Biochemistry Introduction We have just seen how photoexcitation of the non-reaction center chlorophyll turns that molecule into a good reducing agent, which transfers its electron to the nearest excited state level of the reaction center chlorophyll. If you count both steps together, the non-reaction center chlorophyll gets "photooxidized", in the process producing the "strong" oxidizing agent which is the positively charged chlorophyll derivative. The extra electron passed onto the second molecule will eventually be passed on to NADP+ to produce NADPH. These reactions occur in the presence of light and hence are called the light reactions. The light reactions of photosynthesis in green plants are shown in Figure \(1\), along with the standard reduction potentials of the participants, the Z scheme. The combined processes of PSII and PSI resemble a "Z" scheme (rotate in your mind the standard reduction potential figure 90 degrees clockwise). In an organization reminiscent of electron transport in mitochondria, water is oxidized by photosystem II (PSII). Electrons from water are moved through PSII to a mobile, hydrophobic molecule, plastoquinone (PQ) to form its reduced form, PQH2. Another photosystem, photosystem I (PS1), is next in the electron transport pathway. It takes electrons from another reduced mobile carrier of electrons, plastocyanin (PCred) to ferredoxin, which becomes a strong reducing agent. Ferredoxin is a protein with an Fe-S cluster (Fe-S-Fe-S in a 4-membered ring, with 2 additional cysteine residues coordinating each Fe). It ultimately passes its electrons along to NADP+ to form NADPH. Note the complexes that produce a transmembrane proton gradient. In contrast to mitochondria, the lumen (as compared to the mitochondrial matrix) becomes more acidic than the stroma. Protons then can move down a concentration gradient through the C0C1ATPase to produce ATP required for the reductive biosynthesis of glucose. Figure \(2\) a more detailed view of the molecular players in the light reaction. Photosystem II PSII has a complicated structure with many polypeptide chains, lots of chlorophylls, and Mn, Ca, and Fe ions. A Mn cluster, called the oxygen-evolving complex, OEC (also called the OEX) is directly involved in the oxidation of water. Two key homologous 32 KD protein subunits, D1 and D2, in PSII are transmembrane proteins and are at the heart of the PSII complex. It has been said of PSII that "Of all the biochemical inventions in the history of life, the machinery to oxidize water — photosystem II — using sunlight is surely one of the grandest." (Sessions, A. et al, Current Biology 19 (2009) The net reaction carried out by PSII is the oxidation of water and reduction of plastoquinone. 2PQ + 2H2O → 2PQH2 + O2 (g) The oxidation number of oxygen in water is -2 and 0 in O2, so this is a loss of electrons or oxidation of the water. Note that water is not converted to 2H2 + O2, as in the electrolysis of water. Rather the Hs are removed from the water as protons in the lumen of the chloroplast, since the part of PSII that oxides water is near the lumenal end of the transmembrane complex. Protons are required to protonate the reduced (anionic) form of plastoquinone to form PQH2, an activity of PSII found closer to the stroma, derive from the stroma. That being said, researchers actively trying to develop a photosynthetic scheme or mimic that does produce H2 for use as a clean and essentially boundless fuel source to replace climate-warming fossil fuels. A quick look at standard reduction potentials (SRP) shows that the passing of electrons from water (dioxygen SRP = +0.816 V) to plastoquinone (approx SRP of 0.11 ) is not thermodynamically favored. The process is driven thermodynamically by the energy of the absorbed photons. The crystal structure of PSII from a photosynthetic cyanobacterium consists of 17 polypeptide subunits with metal and pigment cofactors and over 45,000 atoms. Of particular interest is the P680 chlorophyll reaction center, which consists of four monomeric chlorophylls adjacent to a key Tyr 161 side chain. When H2O gets oxidized to form dioxygen, 4 electrons must be removed by photoactivated P680. In PSII, this process occurs in 4, single electron steps, with the electrons first being transferred to the oxygen-evolving complex. The electrons passed through the Mn complex are delivered to P680 by a photoactive Tyr 161 (Tyr Z or YZ) free radical. Five discrete intermediates of the OEC, S0-S4, are suggested from the experimental data and are consistent with the Kok cycle, which we will discuss below. These were postulated from experiments in which spinach chloroplasts were illuminated with short light pulses. A pattern of dioxygen release was noted that repeated after 4 flashes. Ultimately, light absorption by P680 forms excited state P680*, which donates an electron to pheophytin, which passes them to quinones. Hence P680 gets photooxidized as it forms the cationic P680+. This then removes an electron from Tyr 161 (YZ) producing the tyrosine radical cation, Tyr 161.+. Given its positive charge, its reactive nature as a free radical, and its proximity to Mn ions in the OEC, it pulls an electron from a Mn ion in the OEC. This process repeats itself 4 times for the oxidation of two H2Os, which injects 4 electrons back into the OEC to return to the basal state. The mechanism is very complicated and still not fully understood. It is perhaps easiest to think about the mechanism involving a series of sequential electron and proton transfer and their accompanying change in charge and redox states. Most biochemistry students have a limited understanding of transition state complexes and chemistry but even the experts struggle with the mechanism. In summary, for PSII in plants: 1. a pair of chlorophylls (P680) in the D subunits absorb light (maximum absorbance around 680 nm) and reach an excited state 2. electron transfer from P680 to a nearby chlorophyll with a lower energy level for the excited state electron occurs, which produces an anionic chlorophyll. This chlorophyll has 2 H+ ions in the chlorophyll instead of Mg2+ (again note the charge balance). After electron transfer, P680 now becomes the cation P680+. 3. This "anionic" chlorophyll transfers an electron to oxidized plastoquinone. 4. The P680+, a strong oxidizing agent, removes one electron from an adjacent Tyr 161 to reform P680 and the radical cation Tyr 161.+. Its proximity to the OEC complex leads to it removing an electron from the OEC, making it a more potent oxidizing agent 5. This process repeats a total of 4 times to fully oxidize two water molecules to produce 1 O2 with the 4 electrons removed from 2 glasses of water added back to the metal centers of the OEC. This suggests that there are 5 states of the OEC, an initial state, which we will call S0, and four other states (S1, S2, S3, and S4). S1 forms after the removal of one electron from the OEC by the adjacent radical cation Tyr 161.+ (formed after absorption of one photon). S2, S3, and S4 are sequentially formed after the removal of one electron by a newly regenerated Tyr 161.+ after another round of photoexcitation. S4 then returns to its original state, S0. This series of reactions is called the Kok cycle, which is shown in Figure \(3\). There is no structural information given in the above figure. What is shown instead are possible and consistent oxidation numbers of the four Mnn+ ions in the OEC that are consistent with charge balance and the changes in the oxidation number (-2) of the oxygen atom in water as it progresses to O2 with an oxidation number of 0. The Mn ion states in the Kok diagram denote different discrete oxidation states where n is the number of oxidative “equivalents” stored in the OEC during cycle progression. Think of the OEC as the key catalyst, which will interact with substrate H2O molecules. We start with the S0 state and must return to it in the full cycle. Remember that when O2 acts as an oxidizing agent in combustion reactions, it forms 2H2O. That requires the addition of four electrons. If done sequentially, the oxygen intermediates include superoxide, peroxide, and oxide, the latter of which when protonated is water. Hence two waters and four cycles are required to remove the four electrons required to produce dioxygen. Intermediate but transient oxygen states are also presumably important in this mechanism. A similar mechanism is found in PSI, except plastocyanin, not dioxygen is oxidized, with electrons moved to ferredoxin. This is likewise a difficult process since the reduction potential for oxidized plastocyanin (the form that can act as a reducing agent) is +0.37 while for ferredoxin it is -0.75. This transfer of electrons is an uphill thermodynamic battle since the more positive the standard reduction potential, the better the oxidizing agent and the more likely the agent becomes reduced. What drives this uphill flow of electrons. Of course, it is the energy input from photon. We won't go into any more detail about PSI since it is very similar to PSII but of course, does not have the OEC. The Oxygen Evolving Complex - OEC Even though this is not a bioinorganic textbook, we must move past the "simple" Kok cycle diagram and look at the actual structure of the minicatalyst, the OEC, and the protein and water (substrate) environment around it to understand the mechanism. The mechanism of the OEC is still not fully understood. It's experimentally difficult to unravel given its complexity as the intermediates are very labile and the x-ray-induced transient alterations in the structure of OEC complicate matters more. Paradoxically it is quite simple in overall terms. Here is the essential reaction: 2H2O + 4 photons → 4 H+(lumen) + 4 e- + O2. The crystal structure of PS2 from T. vulcanus has significantly improved our understanding of the OEC and electron flow on water oxidation. We will concentrate on developing an understanding of the amazing Photosystem II from Thermosynechococcus vulcanus, a cyanobacterium (19 subunits with 35 chlorophylls, two pheophytins, 11 beta carotenes, 2 plastoquinones, 2 heme irons, 1 non-heme iron, 4 Mn ions, 3-4 Ca ions, 3 Cl ions, 1 carbonate ion, and around 2800 water molecules). Nature has appeared to evolve a single gene for the central protein in PSII that binds the OEC. The cluster, Mn4CaO5, appears identical in all photosynthetic organisms and is shown below. Researchers were surprised to find that the Ca ion was an integral part of the basic geometric “framework” of the OEC instead of a Mn which was found to be “dangling” from the basic geometric framework. A detailed structure of the OEC from T. vulcanus is shown in Figure \(4\). Note the basic structure is a distorted cube the metal ions at every other corner separated by oxides. Again, it was a surprise that not all of the 4 Mn2+ ions were in the cubic structure. Note one "dangling" Mn2+ with the other last metal site in the distorted cube occupied by Ca2+. Four oxygens (presumably from waters) are shown interacting with MN4 and CA1. It is very difficult to visualize this structure correctly from a 2D figure. Figure \(5\) shows an interactive iCn3D model of the OEC with bound water of photosystem II from Thermostichus vulcanus (3WU2) which should help in visualizing this structure. (very long load time!) Figure \(5\): OEX with bound water of photosystem II from Thermostichus vulcanus (3WU2). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...F8vwgkDXV6C1S6. (very long load time) The shape outlined by O5-CA-O1-MN1 and MN3-O2-MN2-O3 is similar to cubane (shown above right) Now let's zoom out and view some of the amino acid side chains that interact with the OEC from T. vulcanus. These are shown in Figure \(6\). Figure \(6\): OEC and surrounding amino acids from T. vulcanus The coordination number for all the Mn ions (including those interacting with water) is identical. Note the proximity of Tyr 161 which is involved in electron removal from the OEC after it becomes the radical cation Tyr 161.+ in the primary photooxidation event. Figure \(7\) shows an interactive iCn3D model of the OEX with surrounding amino acids and water in photosystem II from Thermostichus vulcanus (3WU2). (long load time) presented to once again help you better understand the 2D structure shown above. Figure \(7\): OEX with surrounding amino acids and water in photosystem II from Thermostichus vulcanus (3WU2). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...bP6ttbaWUmQAg7 The OEC can be thought of as a distorted cubane with a MN3-O4-MN4-O5 back. Bonds to O5 are longer than the other bonds which suggest they are weaker than the other metal-oxygen bonds. This could suggest that it may not be an oxo (O2-) ligand but another variant such as OH- (lower charge), which may imply involvement in the splitting of dioxygen in the reaction mechanism. From a mechanistic perspective, an O-O bond must form between two waters. Both sets of bound "waters" (purple spheres in Figure 4 and red spheres in Figure 5 shown without Hs attached if they are present) are close to O5. It is important to remember that electrons removed from the metal ions in the OEC by Tyr 161.+ must be restored to the OEC to allow the catalytic cycle to continue. These electrons come from the waters that get oxidized. Such a reversible loss and gain of electrons most readily occur from the transition state Mn ions, which you all remember from introductory chemistry have multiple oxidation states. To bring back introductory chemistry again, we present the standard reduction potentials of different Mn ions in Table \(1\) below. Reduction reaction Standard Reduction Potential Mn2+ (aq) + 2 e-→ Mn (s) -1.185 MnO4- (aq) + 2 H2O (l) + 3 e- → MnO2 (s) + 4 OH- +0.595 MnO2(s) + 4H+ + e- → Mn3+ + 2H2O +0.95 MnO2(s) + 4H+ + 2e- → Mn2+ + 2H2O +1.23 MnO4- (aq) + 8 H+ (aq) + 5 e- → Mn2+ (aq) + 4 H2O (l) 1.507 MnO4- (aq) + 4 H+ (aq) + 3 e- → MnO2 (s) + 2 H2O (l) 1.679 HMnO4- + 3H+ + 2e- → MnO2(s) + 2H2O +2.09 O2(g) + 4H+ + 4e- → 2H2O +1.229 +1.229 Table \(1\): Standard reduction potentials for Mn ions compared to O2. You should be able to determine the oxidation number of the Mn ion in each compound. Based on standard reduction potentials, which oxidation states might be sufficient for the oxidation of H2O in the OEC? How does this translate into structural/chemical changes in the OEC? Figure \(8\) provides a recent mechanism consistent with each of the Kok states (S0-S4). The proposed change in redox state for each Mn ion is illustrated with Mn (III) ions in purple and Mn (IV) ions in yellow. Note the change in the oxidation state of the 4 Mn ions from (III, IV, III, and III) in S0 to (IV, IV, IV, and IV) for all of them in S3 and S4. A flip in the side chain of E180 in S2 allows the binding of Mn1 through an oxy link to Ca. ​​ There are many possible different forms of oxygens in the structure including waters, oxides (bridging oxos and possibly terminal oxides), and hydroxides, and the exact form at some sites are still a bit uncertain. Note also the elegance of having a Mn4 cluster to catalyze the 4 electron oxidation of 2 water through the loss of 4 electrons. Also, the redox state change in the Mn ions is different than the one shown in the Kok diagram in Figure 3. In addition, the final O2-producing step going from S4 → S0 is still uncertain. The above mechanism is based on x-ray structures of intermediates and quantum calculations. In it, S4 has an Mn(IV)O. that bonds with the bridging O5 to form O2. Waters As water is a reactant in PSII, there must be water channels leading to the OEC that provide a way for water to enter and for protons to be removed and directed to the lumen to develop a proton gradient. Another rendering of the Kok cycle, the position of the OEC in PSII on the luminal side of the membrane, and the presence of water channels (Cl1, O4, O1) and the Yz network, which connect Tyr 161 (Yz) to the lumen, are shown in Figure \(9\). A more detailed representation of water channels and the Yz networks is shown in Figure \(10\). These structures show that there is no direct water pathway from across the OEC and that all channels restrict water movement to some degree. The O4 and Cl1 channels are narrower than the O1 channel so water in those is less mobile. In the O4 channels, waters 50-53 are near charged groups and are close to a major bottleneck (residues D1-N338, D2-N350, and CP43-P334, -L334). Based on the x-ray structures and molecular dynamic simulations, it appears that the O1 channels allow access of water to the OEC. The Cl1 channel A, which is more rigid, may be involved in H+ transfer during S2 → S3. The Last Step: Electron Transfer to Plastoquinone We are almost ready for the next section in which we will present the flow of electrons away from PSII through mobile electron carriers, leading to the synthesis of NADPH for the reductive biosynthesis of carbohydrates. Before we leave PSII, let's look at what happens to the radical anion P680-, also known as PheA- (pheophytin A) or PheAD1, the reaction center chlorophyll without a central Mg2+ ion) which received an electron from photooxidation of the reaction center P680, as summarized again in Figure \(11\). Pheophytin A- passes its electron to plastoquinone A (in PSII). which passes it on to the lipophilic mobile electron carrier in the thylakoid membrane plastoquinone. It is similar to the mobile electron carrier in mitochondrial electron transport, ubiquinone. Ultimately these are passed to NADP+ to form NADPH for reductive biosynthesis. Figure \(12\) shows an interactive iCn3D model highlighting just the OEX, pheophytin A (PHO), and plastoquinone A (PL9) in photosystem II from Thermostichus vulcanus (3WU2). (long load time). Figure \(12\): the OEX, pheophytin A (PHO), and plastoquinone A (PL9) in photosystem II from Thermostichus vulcanus (3WU2). (long load time). (Copyright; author via source). Click the image for a popup or use this external link:https://structure.ncbi.nlm.nih.gov/i...H7Mycbx1PsFbt6 The OEX (OEC) is shown in spacefill, CPK colors, PHO is shown in spacefill magenta, and PL9 in spacefill cyan. Note the proximity of PHO and PL9 for easy electron transfer to plastoquinone A. Given the number and proximity of high-energy reactive species in the reaction center, it should not be surprising that side reactions can occur. These would decrease the efficiency of light energy transduction and also could damage molecular components of PSII (and similarly PSI) Figure \(13\)s shows a standard reduction potential diagram for the P680 - Pheophytin A - PlastoQuinone A triad (abbreviated P Phe Q) in PSII. Safe routes for charge recombination between P+ and QA are indicated in blue, the damaging route producing 1O2 in red, and the radiative pathway in green. Stabilization of the P+PheoQA state helps prevent reverse electron flow to form P+PheoQA and subsequent charge recombination to form PPheoQA. For clarity, the details of the additional electron transfer steps, including the oxidation of water and the reduction of plastoquinone to plastoquinol by PSII, collectively termed photosynthesis, are omitted. Abbreviations: P, primary electron donor of PSII; Pheo, pheophytin electron acceptor; QA, primary plastoquinone electron acceptor; 1O2, singlet oxygen; 3O2, triplet oxygen; 3P, triplet excited state of P.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/20%3A_Photosynthesis_and_Carbohydrate_Synthesis_in_Plants/20.02%3A_The_Kok_Cycle_and_Oxygen_Evolving_Complex_of_Ph.txt

Search Fundamentals of Biochemistry Introduction In the previous sections, we studied light absorption by chlorophylls, the transfer of energy to the reaction center of photosystem II, the oxidation of H2O by the oxygen-evolving complex (OEC), and the transfer of electrons from these events to the lipophilic carrier of electrons, plastoquinone. Now we are ready to see how the process continues as electrons are passed on from reduced plastoquinone to the cytochrome b6f complex, through photosystem I (which has no OEC) and on to the terminal electron acceptor NADP+, which forms NADPH. This is used for reductive biosynthesis of glucose after fixation of atmospheric CO2 by ribulose bisphosphate carboxylase (RuBisCo). As we saw in mitochondrial electron transport, this passage of electrons is accompanied by the movement of protons from the lumen to the stroma with the ultimate collapse back into the lumen through a rotatory ATP synthase to form the ATP required for reductive biosynthesis. Figure \(1\) reviews again the light reactions of photosynthesis. Cytochrome b6f This complex moves electrons from the mobile lipophilic electron carrier reduced plastoquinol (PQH2), an isoprenoid quinone, to the mobile Cu-containing protein plastocyanin, which plays an analogous role to the mobile protein carrier in mitochondrial electron transport, cytochrome C. It catalyzes the rate-limiting step in electron transport in the light reactions. Figure \(2\) shows an interactive iCn3D model of the spinach plastocyanin (1AG6) Figure \(2\): Spinach plastocyanin (1AG6). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...DwSDDrYMUb9Xn7 The complex that mediates electron transfer between reduced plastoquinone and the protein plastocyanin, cytochrome b6f, is centrally positioned between the two photosystems. In addition, it moves 2 H+s from the stroma into the lumen. These are joined by 2 H+s from the oxidation of water by the OEC (on the luminal side of PSII) to create a transmembrane proton gradient, which will power ATP synthesis. Electrons transfer with cytochrome b6f takes place through the quinol (Q) cycle in a similar fashion to Complex III in mitochondrial electron transport so we won't go into much detail here. Figure \(3\) shows a summary diagram with electron and proton flow. Figure \(4\) shows another version of the Q cycle as an alternative representation. The left box shows the b6f complex transfers two protons (green arrows) per electron transferred (blue arrows) along high (Fe2S2 cluster, cytochrome f) and low potential chains (bl, bh, ci hemes) as well as Quinol (QH2) oxidation at Qo site, Quinone (Q) reduction at Qi site. The right structure depicts haems b (purple), ci and f (red), Fe2S2 cluster (yellow and orange ball-and-stick model), cytochrome b6 (cyan), subunit IV (blue), Rieske subunit (yellow), cytochrome f (red), PetG, L, M and N subunits (green). Magnification of Qi site comprising bh and ci haems. Both cytochrome bc1 and cytb6f are dimeric complexes, with 2 Fe2S2 clusters, two cytochrome bs, and a cytochrome c. The cytb6f complex also has 9-cis β-carotene and additional c1 heme. The electrons move from PQH2 through the complex in a similar fashion as in the bc1 complex. PQH2 is oxidized at the Qp site with a bifurcation of electrons: • one electron moves through the high potential Fe2S2 center and cyt f pathway (bottom left in pathway diagram of Figure 3), with the electron moving to the soluble peripheral protein plastocyanin and one to photosystem I. • the other moves through the low potential bl, bh, ci hemes pathway (top left in pathway diagram of Figure 3), with the electron moving to a plastoquinone at the Qn site near the stroma. A second round of oxidation of PQH2 at the Qp site eventually donates another electron to a plastoquinone-. which regenerates PQH2 after addition of 2 H+ from the stroma. Having two successive oxidations of PQH2 leads to twice the number of protons moving into the lumen. In the process 2 H+ move into the lumen. Cytochrome b6f also is a redox sensor of the status of the plastoquinol/plastoquinone pool. Figure \(5\) shows an interactive iCn3D model of the spinach cytochrome b6f complex (6RQF) Figure \(5\): Spinach cytochrome b6f complex (6RQF). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...tgUhjekB4Tndw6 The region away from the red leaflet represents the lumen side of the complex. The FeS cluster and heme C are in the luminal domains. The coloring scheme is as follows: • cytochrome bcf subunits 4, 5, 6, 7, and 8: light gray • cytochrome b6: dark gray • cytochrome b6 FeS subunit: plum • cytochrome F: cyan • chlorophylls: orange spacefill, labeled • FeS clusters: CPK spacefill, labeled • HEC - Heme C: CPK spacefill, labeled • heme - porphoryin IX containing Fe: yellow spacefill Cytochrome b6f is the rate-limiting step for electron flow but what it's role in regulating the photosynthetic pathway (light reaction plus the dark reaction of carbon metabolism? Data suggest that the complex regulates electron transport in low light conditions but effects a switch to carbon metabolism under saturating light. Johnson and Berry have analyzed electron flow with carbon metabolism in a fashion analogous to transistors in a circuit board, as illustrated in Figure \(6\). They define a transistor as a regulated circuit element that uses variable conductance to control current flow. The linear flow of electrons from water to reductant is viewed as a light-driven current that is under the control of many regulatory feedbacks stemming from carbon metabolism. In limiting light, Cyt b6f presents maximal conductance to flow, and feedback from carbon metabolism adjusts the excitation of PS I and PS II in such a way as to balance the relative rates of linear and cyclic electron flow to the NADPH, Fd, and ATP requirements of the sinks. When the light becomes saturating, feedback from carbon metabolism also decreases the apparent conductance of Cyt b6f, controlling the linear flow of electrons through the plastoquinone pool and the associated flow of protons into the thylakoid lumen. In this way, the regulation of Cyt b6f simultaneously permits efficient photosynthesis and protects the system from photodamage. This model is organized around the idea that the distribution of excitation between PS II and PS I is regulated in such a way as to minimize losses of absorbed light and maximize potential electron transport through Cyt b6f. The expression for the potential electron transport rate has the form of a Michaelis-Menten expression for a single substrate (i.e., light), but describes the kinetic behavior of the entire electron transport chain (i.e., including both photochemical and biochemical reactions). It predicts that electron transport has a hyperbolic dependence on irradiance, with the maximum efficiency realized at the limit where absorbed irradiance goes to zero and the maximum speed realized at the limit where absorbed irradiance is infinite. The trade-off between the speed and efficiency of potential electron transport is driven by the need for the supplies of reduced plastoquinone and oxidized plastocyanin to be balanced to sustain Cyt b6f turnover at the maximum catalytic rate. This causes progressive closure of the PS II and PS I reaction centers, with PS II accumulating in a reduced state and PS I in an oxidized state. As the excitation pressure on PS II and PS I increases, the closure of the reaction centers causes the photochemical yields of PS II and PS I as well as the absorbed quantum yield to decrease as the potential electron flow through Cyt b6f and the potential photosynthetic rate increase" The key prediction of the expression for the potential electron transport rate is that the maximum activity of Cyt b6f defines the upper limit for the theoretical maximum speed of electron transport. The expressions describing feedback control over Cyt b6f activity are based on the idea that Cyt b6f functions like a transistor, i.e., a component of an electrical circuit that uses variable conductance to control current. The fact that Cyt b6f can modulate its conductance to linear electron flow within milliseconds of a perturbation in light suggests that photosynthetic control is the first line of defense against overexcitation, protecting the acceptor side of PS I from being flooded with highly reduced intermediates. In response to a sustained increase in light, the induction of photosynthetic control is followed by the induction of PQN. As NPQ alleviates the electron overpressure in the PQ pool, photosynthetic control progressively relaxes. The two forms of regulation gradually settle to a steady state at the new light intensity. This interaction seems to allow electron transport to proceed at the Cyt b6f-limited rate under low light intensities, and then smoothly switch to the Rubisco-limited rate once the light intensity is high enough to become saturating. It also seems to allow photosynthesis to operate safely and efficiently in a wide range of biochemical milieus, from those characteristic of natural variation in photosynthetic capacity (with balanced electron transport and carbon metabolism) to those characteristic of genetic manipulations (with imbalances in electron transport and carbon metabolism). In this framework, the excitation balance of PS II and PS I and the maximum activities of Cyt b6f and Rubisco emerge as key limits on system dynamics. For example, the trade-off between the speed and efficiency of electron transport is shown to be controlled by the excitation balance of PS II and PS I and the maximum activity of Cyt b6f. The development of PQN is shown to be controlled by the excitation balance of PS II and PS I and the demand for linear electron flow (LEF) through the light reactions and circular electron flow (CEF) around PSI. The onset of photosynthetic control is shown to be dependent on the maximum activities of Cyt b6f and Rubisco. Plant photosystem I-LHCI super-complex If the goal of the photosystem complexes is to transduce light energy delivered by photons into electrons that can be used for reductive biosynthesis of glucose from atmospheric carbon capture of CO2, then PSI is quite amazing. It has a "quantum efficiency" close to 1 which implies that one absorbed photon produces 1 electron that can be used to reduce NADP+. This happens since the transfer of photon energy to other molecules in PSI is so quick compared to nonradiative decay processes for the excited state chlorophylls. The same process for excitation and electron (charge) transfer that we saw in PSII occurs in PSI, with the chlorophyll in the reaction center involved in charge transfers. The light-absorbing molecules of PSI enable light at the far red of the spectra to be absorbed. The complex has 16 proteins, 155 chlorophyll, and 35 carotene derivatives. As with PSII, there is a core complex that is similar to cyanobacterial PSI. The supercomplex has in addition light-harvesting complex I proteins around one side of the complex (looking down on it) that have four LHCI proteins (Lhca 1-4) which allow for more absorption of life. They have 57 chlorophylls and 13 carotene derivatives. The apoproteins that bind the chlorophylls, including the LHCI proteins, have similar but slightly different topologies, allowing for tuning of the absorbance spectra of the bound chromophores. Seven of the chlorophylls when bound have local environments that allow absorption of far red light. These are not found in PSII. the far red light would be more abundant in the low parts of the plant canopies as photons of lower wavelength would be more filtered out by upper leaves in the canopy. Because of the complexity of PSI, we will offer several different iCn3D models to illustrate different features of the photosystem I-LHCI super-complex. As with PSII, PSI has a core with a surrounding light-harvesting complex (LHCI). Figure \(7\) shows an interactive iCn3D model of the Plant (pea) photosystem I-LHCI super-complex (4XK8) Figure \(7\): Plant photosystem I-LHCI super-complex (4XK8). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...uWBzFrVBUDHHQ6 (long load time) Again the red leaflet represents the lumen side of the complex. Gray represents the chlorophyll and other lipids molecules in the complex. Here is the same complex but just the nonprotein components Figure \(8\) shows an interactive iCn3D model of the lipid and FeS components of Plant photosystem I-LHCI super-complex (4XK8) Figure \(8\): Lipid and FeS components of plant photosystem I-LHCI super-complex (4XK8). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?VtoCnxLRHQ8RPdESA (long load time) Again the red leaflet represents the lumen side of the complex. Gray represents the chlorophyll and other lipids molecules in the complex. Rotate the image to see how the chlorophyll and other lipids encircle the membrane protein components. Figure \(9\) shows an interactive iCn3D model of the ApoA1 protein with surrounding chlorophyll and other components from plant photosystem I-LHCI super-complex (4XK8) Figure \(9\): ApoA1 protein with surrounding chlorophyll and other components from plant photosystem I-LHCI super-complex (4XK8). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...L6tsJ3oMDZ2YQA (long load time) ApoA1 is shown in cartoon and colored according to hydrophobicity. The chlorophylls, carotenes, and other components within 5 Å are shown in red. Rotate the image to see how the chlorophyll and other lipids encircle the membrane protein components. Figure \(10\) shows an interactive iCn3D model of the Plant (pea) photosystem I-LHCI super-complex highlighting LHCIs (4XK8) Figure \(10\): Plant (pea) photosystem I-LHCI super-complex highlighting LHCIs (4XK8). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...9vHynkRTLxAct7 (long load time) The LHCI subunits (Lhca 1-4) are shown in alternating magenta and cyan cartoon structure surrounding one side of the core complex. The view in the model above is a top-down view. We discussed in Chapter 20.1 that PSI absorbance is tilted toward the red end of the spectra, with the effective absorbance ratio over a broad wavelength range for the two photosystems, PSI/(PSI+PSII) deviating from around 0.5 at the red/far red end of the spectrum (670-730 nm), where the ratio is close to 1. When exposed to far-red light, the systems move to state I. In this state, the major mobile antenna proteins (LHCIIs) move to PSII to restore a "photoabsorption" balance. When exposed to light depleted in the high end of visual spectra, the system moves to state II, in which mobile LHCIIs move to PSI. The location of LHCs is regulated by the phosphorylation of LHCs controlled by the levels of plastoquinone, which makes great biological sense. When the concentration of plastoquinones in the reduced state, PQH2 (plastoquinol), is high, it would be optimal to increase the activity of PSI to relieve the high concentration of the substrate for cyto b6f and shift the system to higher PSI activity and continue electron flow. This regulation is mediated by the phosphorylation of LHCs by LHCII kinase (a Ser/Thr kinase), which is activated by high reduced PQH2 concentrations. Phosphorylation of LHCs leads to their movement from PSII to PSI. When the oxidized form for plastoquinone is high, LHCII kinase is inactivated by dephosphorylation, causing the mobile LHC to move back to PSII to increase output (PQH2) from PSII. These events occur in low light. In high-light conditions, when the system is functioning at a high level, the LHCII kinase is inhibited by stromal thioredoxin. Specifically, the phosphorylation status of Lhcb1 and Lhcb2 in LHCII homo- or heterotrimers determine the movement between PSII and PSI. The chloroplastic serine/threonine-protein kinase (STT7 also known as STN7) is another LHCII kinase. Chloroplast ATP synthase - CF1FOATPase Finally, let's take a quick look at the ATP synthase in chloroplasts. It is similar in structure and function to mitochondrial FoF1ATPase so we won't spend much time on the mechanism. Like its mitochondrial counterpart, it is a rotary enzyme that transduces the free energy of a proton gradient collapse into chemical energy in the form of ATP. One difference is the rotary enzyme should be regulated by light levels with its activity decreased at night. This is accomplished in higher plants by conversion between a reduced and oxidized state, an effective redox switch, in one subunit (γ) of the ATP synthase. The deactivation is important at night since if the rotary enzyme runs in reverse, ATP hydrolysis would ensue. Figure \(11\) shows an interactive iCn3D model of the reduced R1 state of chloroplast ATP synthase (R1, CF1FO) (6VON). It should look familiar to you given its similarity to mitochondrial ATP synthase. Figure \(11\): Reduced R1 state of chloroplast ATP synthase (R1, CF1FO) (6VON) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?hbnDCm1Kgqgu2k1NA (long load) • alpha - light green • beta - dark green • gamma - yellow • gamma - crimson • epsilon - indigo • b - blue • b' - light blue • a - light pink • c ring - purple • ATP - spacefill, cyan • TENTOXIN (TTX) - space fill gray • ADP - spacefill magenta In the oxidized state, there is a disulfide in the γ subunit which inhibits torsion by stabilizing two β hairpins. This constraint is relieved on reduction and rotation is enhanced. In the reduced structure, tentoxin, an uncompetitive inhibit is present which allowed the structure to be determined. below Yang, JH., Williams, D., Kandiah, E. et al. Structural basis of redox modulation on chloroplast ATP synthase. Commun Biol 3, 482 (2020). https://doi.org/10.1038/s42003-020-01221-8. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/. At night, the enzyme is in the low-activity form. When light becomes available in the day, the pH gradient collapse leads to the release of tightly bound ATP from the oxidized state of the enzyme. PSI can engage in photon-induced electron transfer to ferredoxin. This binds to NADP reductase and leads to NADPH formation. These processes are illustrated in Figure \(12\). Increased reduced ferredoxin leads to the formation of reduced thioredoxin, a small redox protein involved in redox signaling and protection of cells from oxidative stress. A similar process activates the enzyme sedoheptulose-1,7-bisphosphatase (SBPase) found in the dark reaction Calvin cycle of photosynthesis. Its activation is illustrated in Figure \(13\). Reduced thioredoxin also reduces and activates chloroplastic ATP synthase. Figure \(14\) the activities and structures of the ATP synthase (CF1FO) Panel a: For the experiments, purified CF1FO was reconstituted into a liposome (orange) mixed with lipids of phosphatidylcholine and phosphatidic acid. The generated pH gradient across the membrane drove the reconstituted CF1FO to synthesize ATP molecules, which were detected using a luciferin/luciferase assay (green). Val indicates valinomycin. Pane b: The activity of the enzyme is shown in the reduced state (in the presence of dithiothreitol - DTT, blue curve), oxidized state (in the presence of iodosobenzoate - IBZ, orange curve, and with no additive (green curve). Note that the activity is reduced by about 80% in the oxidized state. Panel c: The images are from Cryo-EM density maps of the oxidized and reduced forms of the CF1FO. The color codes are as follows: α (light green), β (dark green), δ (yellow), bb' (blue and light blue), γ (crimson), ε (indigo), a (light pink), and c ring (purple). R indicates a reduced state, and O indicates an oxidized state. The three-dimensional (3D) reconstructions are categorized into three different rotary states (states 1, 2, and 3). The upper insets are the density maps of the F1 domains. Figure \(15\) shows the details of the conformation changes in the γ subunit between the oxidized and reduced forms. Panel a: Structures of the reduced (light blue) and oxidized (orange) γ subunits. Two β hairpin structures (from γGlu238 to γLeu282) are shown in light green, and the two cysteines of the redox switch are shown in yellow in circular enlarged views. The diagram on right shows the topology of the two β hairpin structures. Panel b Superposition of the reduced and oxidized γ subunits (RMSD 1.016 Å). The two β hairpins are shown in light blue and orange for the reduced and oxidized forms, respectively. Other regions are shown in white. Panel c Interaction networks of the β hairpin 2 and βDELSEED motif. The left and right panels are the reduced (γ subunit in light blue) and oxidized (γ subunit in orange) forms. Light green represents the β hairpin 2, dark green for the β subunit, and yellow for the βDELSEED motif. The distances connecting the residues of the γ coiled-coil (γArg73, γGln76, and γGlu77) with the βGlu412 are labeled. Panel d Interaction of the EDE motif with the γ subunit. The EDE motif (yellow) does not interact with any part of the reduced γ subunit but forms an extensive interaction network with its neighborhood when the γ subunit is oxidized. Figure \(16\) shows a cartoon model illustrating the structures of the overall oxidized and reduced states of ATP synthase Panel a: This shows a cartoon schematic of the redox modulation. The upper and lower models are the reduced and oxidized states, respectively. Color codes are the same as in Fig. 1c and the β hairpin structures of the γ subunit are shown in light green. The two redox states are aligned in the same view. Pane b: At night, no energy input from light is available for the photosynthetic electron transport chain, and thus, no electrochemical potential (ΔΨ) and proton gradient (ΔpH) are generated. The oxidized γ subunit prevents CF1FO from hydrolyzing ATP. During the day, light induces charge separation to generate an electrochemical potential across the membrane. Although the CF1FO begins to synthesize ATP molecules, the γ subunit is still oxidized while ΔΨ is small. The rate of ATP synthesis is not at its maximum. At sunrise, thioredoxin subsequently reduces the γ subunit, fully activating CF1FO. The molecular motor, consisting of the γ-ε central shaft and the c14-ring, is free to rotate at full speed to maximize its ATP synthesis activity. Three ATP molecules per rotation of the c14 ring are produced. At sunset, the membrane becomes de-energized, leading to small ΔΨ and ΔpH, and the ATP hydrolysis starts to take place. To prevent ATP loss from excess ATP hydrolysis, the γ subunit is then oxidized again. This process of light regulation and redox modulation on the CF1FO will cycle daily.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/20%3A_Photosynthesis_and_Carbohydrate_Synthesis_in_Plants/20.03%3A_Plant_Electron_Transport_and_ATP_Synthesis.txt

Search Fundamentals of Biochemistry The source for the organization and some of the text derives from: Sindayigaya and Longhini. https://www.peoi.org/Courses/Courses...chem/biochem18 CC - https://creativecommons.org/licenses...sa/3.0/deed.en Introduction We focused on the light reactions of photosynthesis. Now let's turn our attention to the dark reactions which fix CO2 from the air and reduce it with NADPH produced, along with O2, in the light reactions, to produce carbohydrates. The dark reactions don't just occur in the dark. The term is simply used to differentiate them from the light-driven reactions using PSII and PSI. What is so interesting about plants is they produce fuel from CO2 using photons as a source of energy (they are autotrophs) and also consume the fuels they make, using both anaerobic and aerobic respiration pathways. Their biosynthetic reactions take place mostly in the chloroplast, a type of plastid, which are subcellular organelles with specific functions such as photosynthesis or metabolite synthesis and storage. Plants also can not move to acquire fuel and nutrient molecules. They are subject to a large range of growing conditions (differential light qualities and quantities, temperatures, and rainfall levels). Also, plant cells have cell walls in addition to a cell membrane. A simple cartoon showing the major motifs of photosynthesis is shown in Figure $1$. In this section, we will discuss how CO2 from the atmosphere is "fixed" or "captured" in the formation of the simplest sugars (3 carbon molecules like glyceraldehyde-3-phosphate) in a process called the C3 or Calvin Cycle, which is also called the Calvin–Benson–Bassham (CBB) cycle, or the reductive pentose phosphate cycle (RPP cycle). Plants that use the C3 cycle are logically called C3 plants There are two other major types of carbon capture pathways, the C4 and CAM pathways, which we discuss in the next section. All use a key enzyme, ribulose 1,5-bisphosphate carboxylase (RuBisCo), to covalently fix CO2 into small carbohydrates, 3-phosphoglycerate. RuBisCo is the most abundant protein in the biosphere. Recent estimates suggest that there are about 0.7 gigatons (Gt = 1012 tons) of it, with over 90% in the leaves (about 3% of their weight) of terrestrial plants. It captures about 120 Gt of atmospheric CO2 each year. This enzyme has a second competing enzymatic activity. It is also an oxygenase, which adds to its complexity. That activity captures about 100 Gt of atmospheric O2 each year. In this chapter section, we will give an overview of the C3 pathway and given the importance of RuBisCo, we will focus on it predominately. Along RuBisCo, plants have pathways to take the fixed CO2 to 3C sugars and then a unique pentose pathway which runs in a reductive fashion to ultimately produce the sugar-containing molecules in plants we are most familiar with, sucrose and the glucose polymer starch. Plastids There are several types of these organelles. Photosynthesis occurs in chloroplast which has its own genome, like the mitochondria. Another common type is the amyloplasts, which lack pigmented molecules (i.e. they are colorless) and have no inner membrane. In plants, they are filled with starch. Chloroplasts and amyloplasts can interconvert. Chloroplasts are abundant in green leaves while amyloplasts are predominately found in locations like potato tubers where starch is stored. Light can drive the interconversion of plastids as shown in Figure $2$. The characteristics and plastid interconversion pathways of the plastids are shown by arrows. The transition to a chloroplast is called “Greening” and is identified with the number “1”. This is mainly triggered by light signals from proplastids, etioplasts, leucoplasts, and chromoplasts. Etioplasts can develop from proplastids in dark conditions and this is identified by the number “2”. The number “3” indicates leucoplast development that is triggered by diverse development processes to generate starch, lipid, and protein-enriched sub-types called amyloplasts, elaioplasts, and proteinoplasts, respectively. Mainly during the ripening stage, diverse types of carotenoid crystals were generated within the plastids called chromoplasts from the proplastids, leucoplasts, and chloroplasts and this is identified with the number “4”. Together with etioplast and leucoplast development (2,3), chromoplast development (4) was identified as a “Non-greening” plastid transition. The loss of green color from the chloroplasts is called “De-greening” and is identified with the number “5”, and these chloroplasts are then transited into leucoplast or gerontoplast by developmental regulation or during senescence, respectively. CO2 capture and the C3 Cycle There are in the synthesis of the simplest carbohydrates (3 carbon polyhydroxy- aldehydes and ketones: 1. Carbon capture or fixation phase. We prefer the term carbon capture as this term is now used to describe how the world is seeking new ways (other than planting billions of trees) to "capture" excess CO2 emitted through the use of fossil fuels. In a reaction catalyzed by RuBisCo, atmospheric CO2 ultimately react with a 5-carbon acceptor molecule, ribulose 1,5-bisphosphate (Ru1,5-BP, 6 carbons in total), to form two molecules of 3-phosphoglycerate (3PG). (2, 3C molecules). 2. Reduction phase: 3-phosphoglycerate is reduced to glyceraldehyde-3-phosphate (G3P). Three CO2s are captured on reaction with 3 Ru1,5-BP to form 6 glyceraldehyde-3-phosphates (G3P). These can readily interconvert to the keto form, dihydroxyacetone phosphate (DHAP). 3. Regeneration phase: Five of the six G3Ps (15 Cs) react to form 3 three molecules of ribulose 1,5-bisphosphate (15 C2) to allow the catalytic C3 cycle to continue. The other G3P moves into the stroma, in the form of DHAP where it can be used in gluconeogenesis (reductive biosynthesis) of glucose. This can be converted to polymer starch and also the disaccharide sucrose (which we will discuss in a future session). An overview of the Calvin or C3 cycle is shown below in Figure $3$. The stoichiometry can be confusing until you count the actual number of carbon atoms and realize that the cycle has to run 3 times to enable 3 carbon atoms from 3 CO2 molecules to produce one net glyceraldehyde-3-phosphate (G3P). The G3P leaves the C3 cycle at the low left for glucose synthesis. The conversion of the 5 G3Ps that reform Ru1,5-BP requires ATP as shown below: $\ce{5 glyceraldehyde-3P + 3 ATP → 3 ribulose-1,5-2P + 3 ADP + 2 P_i} \nonumber$ with $\ce{P_i}$ indicating inorganic phosphate. Hence the net equation for 3 turns of the cycle, sufficient to produce 1, G3P is: $\ce{3 CO2 + 6 NADPH + 6 H^{+} + 9 ATP + 5 H2O → glyceraldehyde-3-phosphate (G3P) + 6 NADP^{+} + 9 ADP + 8 P_i } \nonumber$ Even though glucose is not a product of the Calvin cycle, some texts use the following equation to show the stoichiometry to run the C3 cycle enough times (6) to fix 6 $\ce{CO2}$ molecules, enough to make 1 glucose from a simple carbon atom counting perspective. $\ce{6 CO2 + 12 NADPH + 12 H^{+} + 18 ATP + 10 H2O → 2 glyceraldehyde-3-phosphate (G3P) + 12 NADP^{+} + 18 ADP + 16 P_i } \nonumber$ Remember that NADPH and ATP are produced in the light reactions in about the same ratio as they are used in the C3 cycle (2NADPH/3ATPs). The net 8 Pis made as products will react with 8 ADP to regenerate 8 ATP in the light reaction. The 9th Pi is incorporated in a triose-phosphate in the light reaction, so one Pi must be imported from the cytoplasm by an inner membrane triose-phosphate/phosphate translocator, which we will discuss below. In the dark, when ATP and NADPH are not produced, CO2 capture also is inhibited. A more detailed diagram showing the detailed reactions to regenerate ribulose 1,5-bisphosphate (Ru1,5BP) is shown in Figure $4$. Abbreviated reactions for the synthesis of sucrose, glucogenic amino acids, and fatty acids are also shown. You should note the reactions for the conversion of the 6C sugar molecule fructose-6-P (F6P) (glycolytic and gluconeogenic intermediate) to the 5C molecule Ru5P and Ru1-5BP, are analogous to the reactions of the nonoxidative part of the pentose phosphate pathway (PPP) pathway which generates 5C sugars for the synthesis of nucleotides, nucleic acids, and some amino acids. Hence we won't discuss them further. Carbon capture of CO2 into 3-Phosphoglycerate - RuBisCo This key enzyme requires a Mg2+ ion and proceeds through a carbamoylated lysine side chain. The Mg2+ ion orients key side chains. The resulting 6C molecule cleaves into two 2 molecules of 3PG. The RuBisCo family of enzymes can adopt different quaternary structures. A homodimer of two large subunits is the minimum catalytic structure. Often there are in addition two small subunits. The most common form is a 16mer which is found in cyanobacteria, red and brown algae, and all higher plants. A possible mechanism of RuBisCo from Synechococcus elongatus, a unicellular cyanobacterium, is shown in Figure $5$. The carbamoylated lysine side chain is shown in green. Ribulose 1,5- bisphosphate is converted to an enediolate which engages in a nucleophilic attack on the CO2 to form a 6C sugar. Hydroxylation at C-3 of this sugar is followed by aldol cleavage. Ultimately two 3PGs are produced, one of which contains the carbon atom from CO2 (red). Figure $6$ shows an interactive iCn3D model of a single heavy and light chain of ribulose 1,5-bisphosphate carboxylase/oxygenase (RuBisCo) from Synechococcus PCC6301 (1RBL). (long load time) The light chain is shown in cyan and key residues in the heavy chain are shown in CPK-colored sticks and labeled. Bound to the heavy chain is a substrate analog/inhibitor, 2-carboxyarabinitol-1,5-diphosphate. It is produced in plants and in the dark, it inhibits the enzyme. With increasing lights, its concentration decreases. Recent Updates (08/14/23) - Rubisco Reacts with both CO2 and O2 Rubisco is a slow enzyme with a kcat of around 2-10 CO2/sec. In addition, it can bind another substrate, O2, and engage in a competing reaction of photorespiration (oxidation) of ribulose 1,5-bisphosphate to form one molecule of 3-phosphoglycerate (3PG) and one molecule of 2-phosphoglycolate (2PG), as shown (incompletely) in Figure $7$ below. Figure $7$: RuBP conversion by Rubisco through the carboxylase (a) and the oxygenase (b) reactions. Tommasi, I.C. The Mechanism of Rubisco Catalyzed Carboxylation Reaction: Chemical Aspects Involving Acid-Base Chemistry and Functioning of the Molecular Machine. Catalysts 202111, 813. https://doi.org/10.3390/catal11070813.  CC BY) license (https://creativecommons.org/licenses/by/4.0/ Following RuBP (1) enolization, the 2,3-enol(ate) intermediate (2) may react with CO2(a) or O2(b) co-substrates. The carboxylase reaction produces the 2-carboxy-3-keto-arabinitol 1,5-bisphosphate intermediate (3) undergoing protonation to the 2-carboxylic acid before hydration. The C2-C3-scission reaction in C3-gemdiolate (5) is described as occurring in a concerted mechanism upon P1 protonation producing two molecules of 3-phospho-D-glycerate (3PGA, 6). The oxygenase reaction produces 3-phospho-D-glycerate (3PGA,6) and 2-phosphoglycolate (2PG,7) How does the enzyme differentiate the two nonpolar substrates, CO2 and O2?  Given the symmetric arrangement of δand δ- charges in CO2, it has a net 0 dipole, as does O2, so it would not align/orient in a field generated by two poles (+ and -). However, CO2, but not O2, would align in a field generated by four charged poles so it has a quadrupole moment, as shown in Figure $8$ below. Figure $8$:  CO2 aligning in a quadrupole field. (field lines from https://commons.wikimedia.org/wiki/F...quadrupole.svg) The dipole unit is the debye and the quadrupole unit can be expressed in debye.Angstrom. Table $1$ below shows some values for dipole and quadrupole moments for simple gases.  CO2 has the highest quadrupole moment of all these simple gases. Molecule Dipole moment (D) Quadrupole moment (D Å) CO2 0.000 4.30 CH4 0.000 0.02 H2 0.000 0.66 O2 0.000 0.39 CO 0.112 2.5 N2 0.000 1.52 Table $1$: Dipole and Quadrupole moments for some simple gases.  Castro-Muñoz, R., Fíla, V., 2018. Progress on Incorporating Zeolites in Matrimid®5218 Mixed Matrix Membranes towards Gas Separation. Membranes 8, 30.. https://doi.org/10.3390/membranes8020030 The active site has a high electrostatic field gradient in the dimeric form of the enzyme.  Figure $9$ shows an interactive iCn3D model showing the electrostatic potential surface in the active site between two heavy chains of spinach ribulose 1,5-bisphosphate carboxylase/oxygenase (RuBisCo) (8RUC).  It shows that is complex. Figure $9$: Electrostatic potential surface in the active site between two heavy chains of spinach ribulose 1,5-bisphosphate carboxylase/oxygenase (RuBisCo) (8RUC). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...2JG3x4csVnEq67 Two heavy chains are shown in light pink and cyan, and one light chain is shown in gray.  The active site regions are shown as an electrostatic surface potential with blue positive and red negative.  The model has an activated substrate analog, 2-carboxyarabinitol bisphosphate, shown spacefill (hard to see given the electrostatic surface potential).  The residue labeled 201KCX is the carbamate of Lys201. Presumably, the electrostatic field in the active site facilitates through subtle interaction of the quadrupole CO2 compared to O2. The mechanisms that differentiate the binding of CO2 and O2 must also overcome the high intracellular concentration of O2 (around 250 μM) compared to CO2 (7–8 μM in C3 plants and 80 μM in C4 plants). The enhanced affinity for CO2 (about 30-fold) compared to O2 helps overcomes these concentration barriers. No classic "binding pocket" exists for CO2 and O2 so diffusion and binding is likely guided by the electrostatic potential gradients along the diffusion surface. Figure $10$ below the electrostatic potential molecular surface of O2 and CO2 (top), calculated from electron density measurement using quantum mechanics, and the electrostatic surface of their binding pockets Figure $10$: Electrostatic potential molecular surface of O2 and CO2 (top) and the electrostatic surface of their binding pockets.  Tommasi, I.C. et al., ibid. (top) Computed electrostatic potential molecular surfaces of CO2 (left) and O2 (right). The color scheme follows commonly accepted conventions: blue, positive; red, negative. The value of the Qzz component of the quadrupole moment, as calculated by Stec, is −3.239 e a02 for CO2 and −0.232 e a02 for O2.  Note that the ratio of these values is about 14, about equal to the earlier quadrupole moments discussed above with units of debeye.Å) (bottom) a ribbon representation of the catalytic domain with bound gaseous ligands and surfaces colored by the electrostatic potential. O2 (in red) and CO2 (in purple) lie in a positively charged cavity (blue) of the TIM barrel. (Figure from ref. [15], used by permission of PNAS (copyright © 2012)). Both CO2 and O2 are situated in a tiny "cavity" that is blue (positive potential, C-terminal domain) and just above it red (negative potential, N-terminal domain).  The quadrupole moment of CO2 is 10-15x that of O2 which helps explain its higher affinity in the localized electrostatic potential gradients around the gas molecules. Molecular dynamic (MD) simulations show an interaction preference for CO2.  There are many subunit-subunit interfaces and all appear in MD to preferentially interact with CO2, which probably moves from the solvent through large:small subunit interface to the active site.  The CO2 does not localize long at any residues but seems to occupy areas instead.  Since CO2 has no dipole, it locates more closely to small hydrophobic side chain (Ala, Val, Leu, Ile) and the main chain.  CO2 has more interaction in every active site as well as the large:large subunit interface (whose electrostatic potential in the active site is shown above) and in the large:small subunit interface. In efforts to quantitate the preference of CO2 over O2 using MD, investigators found that over many different species of rubisco, the relative distribution of CO2 and O2 to the small and large was on average 1.8 for CO2 and 1.4 for O2 with the number of oxygen bound to either subunit lower. This is true even though CO2 has a lower solvation energy than O2 in water, so additional energy must be spent to differentially desolvate CO2.  The hydrophobic interactions likely promote the movement of CO2 to the active site where electrostatic-based potentials likely favor CO2 binding.  These results suggest that the small subunit acts like a "miniresevoir" for O2 which then diffuse to the large subunit region of the active site.  From a simple thermodynamic perspective, CO2 would be favored to move along the surface and through spaces in the protein guided by transient interactions that be water.  The active site in rubisco is not in a deep pocket but rather in shallow groves near the surface.   Although the enzyme is slow (2-10 CO2/s), it's not much slower than the average enzyme.  The median turnover number kcat (under saturating conditions) of enzymes is about 10 s-1 with most following between 1-100.   Its concentration is very high in chloroplasts, which helps increase the fixing of CO2. The next step: 3-Phosphoglycerate to Glyceraldehyde 3-Phosphate and dihydroxyacetone phosphate The 3PG produced by RuBisCo is converted to the triose glyceraldehyde-3-P (G3P) (which can readily isomerize to dihydroxyacetone phosphate) using typical glycolytic enzymes run in reverse except that NADPH is used as a reductant instead of NADH. In addition, the stromal and cytosolic enzymes derive from different genes. The remaining G3P not used to resynthesize ribulose 1,5BP can be used for the synthesis of starch, sucrose, etc, as illustrated in Figure 4 above. Exchange of trioses and phosphate across the inner membrane The inner chloroplast membrane has a triose-phosphate/phosphate translocator (TPT), an antiporter that brings into the stroma Pi in exchange for a triose phosphate, either dihydroxyacetone phosphate or 3-phosphoglycerate. The importance of this was discussed above. The exported triose can be used for the synthesis of sucrose, which can be transported around the plant as a source of carbon. Trioses within the chloroplast can also be converted to glucose and onto glycogen as the organelle becomes an amyloplast. If the translocator is inhibited, Pi would decrease in the chloroplast, which would decrease ATP and also starch synthesis. The structure of TPT has been determined with the bound ligands, 3-phosphoglycerate and inorganic phosphate, in an occluded conformation from Galdieria sulphuraria, an extremophilic unicellular species of red algae. Figure $11$ shows an interactive iCn3D model of a triose-phosphate/phosphate translocator from the red algae (5Y78). The model shows two monomers, one with red cylindrical alpha helices and spacefill 3-phosphoglycerate. The other subunit is shown in gray with the 3PG in colored sticks and conserved residues (T188, K204, F263, Y339, K362, R363) that make interacts with both Pi and 3PG. There would presumably be an outward- and inward-open conformation that is triggered on ligand binding. Activity Regulation by Light Given their role in photosynthesis, you would expect even the dark reaction enzymes would be regulated by light. Indeed, four C3 cycle enzymes are. They are ribulose 5-phosphate kinase, fructose 1,6-bisphosphatase, sedoheptulose 1,7-bisphosphatase, and glyceraldehyde 3-phosphate dehydrogenase. The regulation is affected by photon-induced disulfide bond formation between two cysteine side chains in the enzymes. When oxidized (disulfide bond form), the enzymes are inactive. Under light conditions, PSII, cyto b6f, and PSI work in electron transport to move electrons from H2O to ferredoxin and onto a small soluble protein thioredoxin which has a disulfide. The enzyme catalyzing this last step is ferredoxin-thioredoxin reductase. On reduction, the disulfide in thioredoxin is cleaved, and the now free sulfhydryls in thioredoxin are used to cleave the disulfide in the 4 enzymes mention above, in a similar fashion to how β-mercaptoethanol in excess can cleave disulfides in proteins. This leads to conformational changes in the four enzymes which activate them. In the absence of light, the process reverses, and the enzymes are inhibited. For fuel at night, plants mobilize starch for energy. A simple mechanism to show how thioredoxin catalyzes disulfide bond reduction in target proteins is shown in Figure $12$. The first enzyme in the oxidative branch of the pentose pathway, glucose 6-phosphate dehydrogenase, uses NADP+ as an oxidizing agent, producing NADPH. In the light, there is lots of NADPH produced from the light reactions of photosynthesis so it makes biological sense that under these conditions, glucose 6-phosphate dehydrogenase activity is inhibited. It is so, also by the cleavage of a critical disulfide bond, but in this case, it results in enzyme inactivation. We saw the role of thioredoxin in the previous chapter section when we discussed the regulation of photosynthesis as well as the ATP synthase of the chloroplast. Figure $13$ shows an interactive iCn3D model showing a comparison of the structures of oxidized (1ERU) and reduced (1ERT) human thioredoxin. Figure $13$: Comparison of the structures of oxidized (1ERU) and reduced (1ERT) human thioredoxin. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...XPVsErV2JWxdA6. The two subunits of thioredoxin, linked by a disulfide are shown in gray. Press the "a" key to toggle between the oxidized form, with Cys32-Cys35 disulfide shown as a yellow stick, and the reduced form with the reduced and separated Cys 32 and Cys 35 shown in colored spheres. Not that the hydrogen covalently attached to the free cysteine side chain does not show in a crystal PDB structure. SUMMARY Photosynthesis in vascular plants takes place in chloroplasts. In the CO2-assimilating reactions (the Calvin cycle), ATP and NADPH are used to reduce CO2 to triose phosphates. These reactions occur in three stages: the fixation reaction itself, catalyzed by Rubisco; reduction of the resulting 3-phosphoglycerate to glyceraldehyde 3-phosphate; and regeneration of ribulose 1,5-bisphosphate from triose phosphates. Rubisco condenses CO2 with ribulose 1,5-bisphosphate, forming an unstable hexose bisphosphate that splits into two molecules of 3-phosphoglycerate. Rubisco is activated by covalent modification (carbamoylation of Lys201) catalyzed by Rubisco activase and is inhibited by a natural transition-state analog, whose concentration rises in the dark and falls during daylight. Stromal isozymes of the glycolytic enzymes catalyze the reduction of 3-phosphoglycerate to glyceraldehyde 3-phosphate; each molecule reduced requires one ATP and one NADPH. The cost of fixing three CO2 into one triose phosphate is nine ATP and six NADPH, which are provided by the light-dependent reactions of photosynthesis. An antiporter in the inner chloroplast membrane exchanges Pi in the cytosol for 3-phosphoglycerate or dihydroxyacetone phosphate produced by CO2 assimilation in the stroma. Oxidation of dihydroxyacetone phosphate in the cytosol generates ATP and NADH, thus moving ATP and reducing equivalents from the chloroplast to the cytosol. Four enzymes of the Calvin cycle are activated indirectly by light and are inactive in the dark so that hexose synthesis does not compete with glycolysis—which is required to provide energy in the dark. Photorespiration - RuBisCo/Oxygenase and the Glycolate Cycle As autotrophs, plants make their fuels. They use that fuel to make ATP to power endergonic reactions like protein synthesis, cell division, etc. As eukaryotic cells, they have mitochondria and can use both aerobic and anaerobic respiration to produce ATP. In the dark, when photons are not present, they carry out mitochondrial aerobic respiration as they break down carbohydrates to CO2 and water, the reverse of photosynthesis. They also use O2 in another process that is driven by light. The same enzyme that captures carbon, RuBisCo, has oxygenase activity. RuBisCo uses O2 in a process called photorespiration, which produces CO2 in a competing reaction. Same enzyme, different substrates! The final products of the reaction with CO2 using RuBisCo are two 3C molecules, 3-phosphoglycerate (3PG). Using O2 as a substrate produces 1 molecule of the 3C 3PG and 1 molecule of a 2C analog, 2-phosphoglycolate (not 2-phosphoglycerate). 2-phosphoglycolate is also named carboxymethylphosphate. About one out of every four turnovers of the enzyme produced this metabolic dead product. Given this non-trivial side reaction, the enzyme should be called ribulose 1,5-bisphosphate carboxylase/oxygenase. You may ask why would such a critical enzyme evolved to a form which is quite inefficient. One explanation is that the enzyme "finished" its evolution before the great oxygenation event when dioxygen rose to the levels we see now (20%). Before that, little oxygen was available to compete with the trace gas CO2, which is around 0.04% of the atmosphere. (Even though CO2 is considered a trace gas, its present concentration is around 420 parts per million (ppm), levels which are warming our planet and which have not been seen for 3 million years, when Arctic forests and camels were present). Compare the KM values (9 μM for CO2 and 350 μM for O2 or 39x higher for O2) for the enzyme and the equilibrium concentrations of the gases in aqueous solution (11 μM for CO2 and 250 μM for O2 or 23x higher for O2). The higher KM for O2 is nearly offset by O2's greater solubility so modern conditions lead to a significant waste of the CO2 capture efficiency of RuBisCo/Oxy. At higher temperatures in a warming world, the equilibrium ratio of solution concentrations of O2/CO2 increases as does the affinity (based crudely on KM values) of CO2. Both of these exacerbate the wasteful oxygenase activity effect. Finally, as CO2 is captured by the enzyme, the ratio of the local concentrations of O2/CO2 also goes up. All of these make the efficiency of RuBisCo worse. Figure $14$ shows a mechanism for the reaction of both CO2 and O2 with RuBisCo/Oxygenase. Note that in contrast to most oxygenases, no cofactor is required for the RuBisCo/Oxygenase. The glycolate pathway The 2-phosphoglycolate (carboxylmethyl phosphate) "waste" product of the oxygenase activity of RuBisCo/Oxygenase is recycled through a complex pathway that is called "photorespiration". It occurs in three different organelles, the chloroplast, the peroxisome, and the mitochondria. Part of the generalized pathway is shown in Figure $15$. Multiplying the stoichiometry represented in the figure by 2 shows that 2 molecules of 2-phosphoglycolate produce 2 molecules of glycine. These get converted to two molecules of serine. We will see the mechanisms for some of these reaction in the chapter on amino acid metabolism. The net reaction is: $\ce{2 Glycine + NAD^{+}+ H2O → serine + CO2 + NH3 + NADH} \nonumber$ The serine is eventually converted to 3-phosphoglycerate, which can be used again in the C3 cycle. Note that CO2 is produced in the glycolate pathways that started with the use of O2 as a substrate by RuBisCo/Oxygenase. Hence the whole system uses O2 and produces one CO2 so the combined reactions are usually called photorespiration. It's not an ideal term since it is wasteful, compared to mitochondrial respiration. Some prefer to call the combined pathway of Rubisco oxygenase and the glycolate pathways the C2 cycle.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/20%3A_Photosynthesis_and_Carbohydrate_Synthesis_in_Plants/20.04%3A_CO_uptake_-_Calvin_Cycle_and_C3_organisms.txt

Search Fundamentals of Biochemistry The source for the organization and some of the text derives from Sindayigaya and Longhini. https://www.peoi.org/Courses/Courses...chem/biochem18 CC - https://creativecommons.org/licenses...sa/3.0/deed.en The C4 Pathway Photorespiration, caused by the oxygenase activity of RuBisCo/Oxygenase and the ensuring glycolate salvage (C2) pathway, significantly diminishes the efficiency of photosynthesis in C3 plants. Rest assured, some plants have found a way around the problem by adding a few steps before RuBisCo. The altered pathway is called the C4 pathway and plants that use it are called C4 plants. The initial steps involve a temporary capture of CO2 in the form of HCO3-, into a 4C, not 3C sugar. Plants that use the C4 pathway include maize, sorghum, sugar cane, and many tropic plants. An overview of the pathways emphasizing the steps that precede RuBisCo is shown in Figure \(1\). Experimental evidence shows that radiolabeled 14CO2 is captured into the 4C molecule, oxaloacetate (OAA), a citric acid cycle and gluconeogenic intermediate, through the enzyme phosphoenolpyruvate carboxylase, which uses HCO3- as a substrate. The reaction takes place in mesophyll cells. OAA can be reduced by NADPH using malate dehydrogenase, or converted to aspartate through a transamination (not shown in the figure). Malate moves in the bundle sheath cell where it is decarboxylated by the NADP-malic enzyme to pyruvate. Pyruvate can move back into the mesophyll cell and be converted to phosphoenolpyruvate (PEP) and then back to OAA by the enzymes pyruvate phosphate dikinase and PEP carboxylase. CO2 from the decarboxylation of malate is delivered as a substrate to RuBisCo. Pyruvate dikinase is used in bacteria, protozoa, C4 plants, and another type, Crassulacean, discussed below. Its non-plant function is to produce ATP, in a fashion similar to pyruvate kinase. The net reaction is: ATP + phosphate + pyruvate = AMP + PPi + H+ + phosphoenolpyruvate Figure \(2\) shows a simplified mechanism for the reaction The net reaction shows that in C4 plants, two molecules are phosphorylated by ATP. One is pyruvate and the other is inorganic phosphate (Pi). Hence the name dikinase. The reverse reaction of ATP synthesis occurs in bacteria and protozoan. There are two phosphorylated intermediates, an Enz-His-P, and an Enz-His-PPi. These served as "activated" phosphate carriers in the phosphotransfer reactions to pyruvate and to Pi, respectively. If PPi is not hydrolyzed to 2Pi, as illustrated in the top left of Figure 2 above, the reaction is fully reversible. In C4 plants there are three reactions 1. Pyr + E-His-P ↔ PEP + E-His 2. E-His + ATP ↔ E-His-PP .AMP (. indicates a noncovalent interaction) 3. E-His-PP .AMP + Pi ↔ E-His-P + AMP + PPi When PPi is hydrolyzed, the net input of ATP to phosphorylated pyruvate is two ATP equivalents. In the next step, PEP is carboxylated in a carbon capture reaction by PEP carboxylase, which as mentioned above used HCO3- as a substrate, not CO2 per se. PEP carboxylase also does not have a competing oxidase activity. The product is malate, which releases locally high "saturating" concentrations of CO2 in the bundle sheath cells, which significantly suppresses the oxygenase activity of RuBisCo. Pyruvate phosphate dikinase undergoes a very large change in a conformational change in domain organization during the catalytic cycle. The two kinase activities are located at different sites in the enzyme. The phosphorylation of Pi occurs in the N-terminal domain, while the phosphorylation of pyruvate is in the C-terminal domain. The center domain that links to the N- and C-terminal domains by associated "tethers" is the site of the catalytic histidine involved in phosphotransfer. A swiveling of domains occurs to allow sequential phosphotransfers. The Pfam domain structure for the protein is shown in Figure \(3\). Figure \(4\) shows an interactive iCn3D model of an AlphaFold predicted model of chloroplast pyruvate, phosphate dikinase from Flaveria brownii (Brown's yellowtops) (Q39734) Figure \(4\): AlphaFold predicted model of chloroplast pyruvate, phosphate dikinase from Flaveria brownii (Brown's yellowtops) (Q39734). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...VB6Aywuc2bLD38 The green represents the N-terminal NBD, the red is the central domain with the catalytic His (side chain shown as CPK-colored spheres and labeled), and the blue the PEP/Pyr PBD). Figure \(5\) illustrates the conformation changes that occur in the central domain (yellow in this figure) and the N-terminal nucleotide-binding domain (NBD, green) in various ligand-bound states. Panel a–c show movement of the central domain (yellow). In (a) it swivels to face the PBD domain (PDB 5JVL/C) and in (c) it faces the NBD domain (TbPPDK (PDB 2X0S)). In (b) it is in an intermediate position. Panels d-f show the movement of the NBD (the three subdomains are depicted by three different greens). Pane (d) is the state without bound nucleotide (PDB 5JVJ/A), while panel (f) shows the fully closed, nucleotide-bound state (PDB 5JVL/A). Panel (e) shows a semi-closed, nucleotide-bound state (PDB 5JVL/C). Minges, A., Ciupka, D., Winkler, C. et al. Structural intermediates and directionality of the swiveling motion of Pyruvate Phosphate Dikinase. Sci Rep 7, 45389 (2017). https://doi.org/10.1038/srep45389. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ Figure \(6\) shows interactions of bound substrates in different conformation states of PPDK. Figure \(6\): Substrate binding sites of FtPPDK. Panel (a) shows the semi-closed state of the PEP binding site (PDB 5JVL/A) with the catalytic H456 (yellow) pointing away from PEP. Panel (b) shows the closed state of the PEP binding site (PDB 5JVL/C) showing interactions between PEP and surrounding residues, including the catalytic H456 (yellow). Panel (c) shows the closed state of the nucleotide-binding site of 5JVL/D occupied with 2'-Br-dAppNHp, a nonhydrolyzable ATP analog. Interacting residues are highlighted. Minges, A et al. ibid. After CO2 is delivered from malate in the bundle sheath cell using RuBisCo, the rest of the reactions are the same as in the C3 pathways. Once CO2 is fixed into 3-phosphoglycerate in the bundle-sheath cells, the other reactions of the Calvin cycle take place exactly as described earlier. Overall the C4 pathways require more ATP. A molecule of PEP is required for each CO2 fixed in the C4 pathway which takes the equivalent of two ATPs. So for each CO2 fixed in the C4 pathway, it takes 5 ATPs compared to 3 ATPs in the C3 pathway. As mentioned above the affinity (estimated from the KM) of CO2 for RuBisCo decrease with increasing temperature, which decreases the energetic efficiency of carbon capture. At higher temperatures (28-30 C), the extra energy cost for the C4 pathway balances out with the extra energy cost for the C3 pathway at higher temperatures. Carbonic Anhydrases We first encountered carbonic anhydrase when we discussed its mechanism in Chapter x.xx. We'll now discuss its function and activity in the C4 pathway in some detail. Given that we are facing a climate crisis due to the increasing levels of CO2 in the atmosphere arising from the burning of fossil fuels, removing CO2 from the atmosphere, a process called for climate purposes carbon sequestration, and understanding the role of carbonic anhydrase in CO2 sequestration becomes even more important. Plants have three different genes for carbonic anhydrase (α, β, and γ). Each can be differentially spliced so there many different isoforms of this protein in plants. They are most abundant in the chloroplast, cytoplasm, and in mitochondria and they have many additional roles outside of fixing CO2 in C4 (and also CAM) plants. In autotrophic (make their own "food") bacteria (such as cyanobacteria, also known as blue-green algae), there are no internal organelles. However, there are carboxysomes, which are protein-bounded vesicles (much like a bacteriophage head), which contain in their internal compartment not nucleic acid but RuBisCo and carbonic anhydrase. The carbonic anhydrase converts HCO3- to CO2 for reaction with RuBisCo. The carboxysome hence concentrates the CO2-producing and fixing enzymes for photosynthesis. Figure \(7\): In C3 plants, CO2 (aq), that is dissolved CO2, is the actual substrate for RuBisCo so available HCO3- is converted to CO2 by carbonic anhydrase. In C4 and CAM plants, CO2 (aq) is first converted to bicarbonate by carbonic anhydrase. HCO3- (aq) is then used as the actual substrate for the "carbon fixation" step. Hence carbonic anhydrase has roles in C3, C4, and CAM plants. All of the carbonic anhydrases have a Zn+2 at the active site. The alpha form, the most prominent in plants, was discovered in erythrocytes and is typically active as a monomer. It has one large 10-strand beta sheet surrounded by 7 alpha helices. The Zn ion is tetrahedrally coordinated by 3 histidine side chains and water. Gamma carbonic anhydrase is a trimer with three active sites at the interface between pairwise monomers with the Zn-coordinating histidine side chains from two different subunits. Beta carbonic anhydrase in plants is typically an octamer of identical subunits. The Zn ion is coordinated by two cysteines, one histidine, and water. The monomer has 4 beta strands in a beta-sheet surrounded by alpha helices. An additional beta-strand is involved in monomer interactions. As the active site is in the interface of two subunits, the functional biological unit is the dimer, but a tetramer and even an octamer are typically formed. The substrate binding groups have a one-to-one correspondence with the functional groups in the alpha-carbonic anhydrase active site, with the corresponding residues being closely superimposable by a mirror plane. Therefore, despite differing folds, alpha- and beta-carbonic anhydrases have converged upon a very similar active site design and are likely to share a common mechanism. Figure \(8\) shows an interactive iCn3D model of beta-carbonic anhydrase from Pisum sativum (pea) with bound acetate (1EKJ). Figure \(8\): Beta-carbonic anhydrase from Pisum sativum (pea) with bound acetate (1EKJ). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...VZTEJPy1hPxK27 For the sake of simplicity, only two subunits (A - gray and B - magenta) of the octamer are shown. Acetate, a proxy for HCO3-2, is shown in spacefill binding between the two subunits in CPK-colored spheres. The side chains involved in Zn2+ binding, Cys 160, His 220, and Cys 223 (unlabeled) are on the A chain. The amino acids that bind acetate are distributed between the A chain (Asp 162, Gly 224, Val 184) and the B chain (Gln 151, Phe 169, and Tyr 205) and are labeled. The same groups are involved in substrate binding in alpha carbonic anhydrase but in a mirror image orientation (but with the normal L-amino acids). Beta carbonic anhydrase has high levels of expression in leaves and is found in chloroplasts, mitochondria, and the cytoplasm. Many plants have it in the cytoplasm and chloroplasts. • β-carbonic anhydrase (βCA) in C3 plants: Most of βCA in leaves is in chloroplasts in mesophyll cells and may comprise up to 2% of leave protein. Yet studies have shown that you can delete the gene for it with minimal effect on the maximal rate of photosynthesis. However, plant development was affected so by interference the enzyme is probably most needed to produce HCO3- for biosynthesis. • β-carbonic anhydrase (βCA) in C4 plants: Most of the βCA is found in the cytoplasm of mesophyll cells. There is catalyzes the first reaction of the C4 pathway, CO2 (aq) to HCO3- (aq). Mitochondrial (βCA) and γCA probably function to fix CO2 arising from oxidative respiration. Crassulacean acid metabolism (CAM) pathway Plants that encounter the chronic stress of low water availability have evolved yet another pathway to adapt to low water conditions. Stomata in C3 plants are open during the day to allow carbon capture from CO2, but they can close when water is limited. This obviously will inhibit plant growth. In the CAM pathway, the stomata stay open at night, allowing carbon capture at a time when water loss through the stomata would be lower. The incoming CO2 is fixed through carbonic anhydrase and then a series of several enzymes to form malic acid, which is transported for storage and use during the light in vacuoles. This CAM pathway is described in Figure \(9\). The proteins and intermediates in the CAM pathways are ALMT9, aluminum-activated malate transporter; CA, carbonic anhydrase; MDH, malate dehydrogenase; OAA, oxaloacetate; ME, malic enzyme (NAD or NADP); P, phosphate; PEPC, phosphoenolpyruvate carboxylase; PEPCK, PEP carboxykinase; PPCK, PEPC kinase; PPDK, pyruvate, phosphate dikinase. During the day, malic acid moves back into the cytoplasm where it is decarboxylated by malic enzyme and releases locally high CO2 concentrations for use by RuBisCo in the C3 cycle. In plants that use CAM, a series of other changes occur, including leaf structure and additional regulatory processes that coordinate metabolic genes. Figure \(10\)s shows more details of the CAM cycle. Thermodynamic comparison between the C3/C4 pathways and the Otto cycle It is interesting to compare the thermodynamic efficiencies of plants and internal combustion engines that are governed by the thermodynamic Otto cycle. In the cycle, chemical energy in the form of gasoline and O2 are converted to thermal energy which is converted into mechanical energy. Both photosynthesis (the conversion of the energy of photons into chemical energy) have limited efficiencies. • In internal combustion engines, power is limited in part by air uptake and by different efficiencies when running at non-constant speeds (like stop-and-go). • In the C3 pathway, efficiency is limited by the oxygenase activity of RuBisCo/Oxygenase (given the much higher concentration of atmospheric O2 compared to CO2) and the ensuring photorespiration pathway. • Water availability also plays a role as the net reaction of photosynthesis and glucose production, in simplified form, is 6CO2 + 6H2O → 6C(H2O6) so in high heat and low humidity the process efficiency decreases Figure \(11\) shows a comparison of three stages, the storage component, the basic cycle, and a concentration mechanism, in photosynthesis and the internal combustion engine (ICE). A concentrating mechanism in C4 plants and turbocharged cars provides concentrated CO2 and oxygen, respectively, to the core cycle (upper row). A storage mechanism in CAM plants allows carbon dioxide to be stored as malic acid at night and then passed to the Calvin cycle during the day, while a storage mechanism in hybrid electric vehicles (HEVs) allows energy to be stored in the battery during braking and then passed to the motor to power the drivetrain in parallel with the engine (bottom row).

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/20%3A_Photosynthesis_and_Carbohydrate_Synthesis_in_Plants/20.05%3A__CO2_uptake_-_C4_and_CAM_Pathways.txt

Search Fundamentals of Biochemistry Now that we have seen how carbon is captured and fixed into 3C trioses, which can be converted to fructose and glucose and their derivative, we can now explore the synthesis of the key plant carbohydrates we all know, sucrose, starch, and cellulose. The source for the organization and some of the text derives from Sindayigaya and Longhini. https://www.peoi.org/Courses/Courses...chem/biochem18. Creative Commons - https://creativecommons.org/licenses...sa/3.0/deed.en Sucrose Synthesis Sucrose is a disaccharide of glucose and fructose with an acetal link between the anomeric carbons to form the nonreducing sugar O-α-D-glucopyranosyl-(1→2)-β-D-fructofuranoside. Its structure is shown in Figure \(1\). Figure \(1\): Sucrose, a disaccharide of glucose and fructose Sucrose can be considered a transport form of carbon, much like ketone bodies are transport forms of fatty acids. As noted above, the link between the sugars is between the anomeric C-1 link of glucose and the anomeric C-2 link of fructose. As such, it is not cleaved by typical carbohydrate-cleaving enzymes like amylases. Also, it doesn't react with proteins like other sugars with free cyclic hemiacetals that can open and form reactive aldehydes. For example, the cyclic monosaccharide glucose, with its anomeric carbon in a readily reversible hemiacetal link, can form covalent bonds to amine groups in proteins such as hemoglobin, which forms the glycosylated version of HbA1c, as shown in Figure \(2\). Figure \(2\): Reactions of cyclic hemiacetal sugars with amines. https://en.wikipedia.org/wiki/Glycated_hemoglobin. Creative Commons Attribution-ShareAlike License 3.0 The reaction proceeds through the formation of a Schiff base followed by a rearrangement. HbA1c is a marker of diabetes. These features of sucrose may explain its choice as a key form of synthesized carbohydrate in plants. Sucrose is synthesized in the cytosol, as shown in Figure \(3\). Figure \(3\): Synthesis of sucrose in plants. We have seen the enzymes that catalyze these reactions through G1P before in the glycolytic and gluconeogenesis pathways. Glucose-1-phosphate is then converted to UDP-glucose, which reacts with fructose-6-phosphate to form sucrose 6-phosphate and UDP, catalyzed by the enzyme sucrose 6-phosphate synthase. The last reaction is the hydrolysis of sucrose 6-phosphate by sucrose 6-phosphatase, which allows the export and transport of sucrose. In the reaction scheme above, G1P is converted to UDP-Glc through the following reaction: G1P + H+ + UTP ↔ UDP-Glc + Pi This reaction is just mildly favored thermodynamically. Before we study the structure of the enzyme, we will first discuss the regulation of the sucrose synthesis pathway and discuss the generic mechanism of glycosyltransferases. Regulation Carbon capture in the light reaction of photosynthesis leads to sucrose (for transport) and starch synthesis. Which product(s) result depends on key regulatory steps. Remember of the 6 trioses formed in the Calvin cycle, 5 are returned into the cycle for the synthesis of ribulose 1,5-bisphosphate, and only 1 is used for the synthesis of sucrose and starch. If too much is removed, the cycle slows. If not enough is used for starch and sucrose synthesis, Pi, which is moved into the stroma from the cytoplasm by an important translocator (see section 20.3) would run low. The key cytosolic regulatory step is catalyzed by fructose 1,6-bisphosphatase (FBPase-1) and also a unique plant enzyme, PPi-dependent phosphofructokinase (PP-PFK-1), that catalyzes the reverse reaction of F6P → F1,6BP, in regulatory steps that are similar to ones found in glycolysis and gluconeogenesis. • fructose 1,6-bisphosphatase (FBPase-1) is inhibited by the allosteric modulator fructose 2,6-bisphosphate (F2,6BP) • PPi-dependent phosphofructokinase (PP-PFK-1) is activated by fructose 2,6-bisphosphate The coordinate regulation of these two enzymes is shown in Figure \(4\). Figure \(4\): Regulation of sucrose synthesis at the formation of fructose-6-phosphate The substrate for the PPi-dependent phosphofructokinase (PP-PFK-1) is PPi, which serves as a phosphodonor. In the plant cytosol, there is no pyrophosphatase to catalyze the cleavage of PPi into Pi. Also note that Fru-2,6-P2 itself is synthesized and degraded by the bifunctional enzyme phosphofructokinase 2/fructose-2,6-bisphosphatase, which we studied before in the regulation of the same step in glycolysis/gluconeogenesis. The levels of F2,6-BP depend on the rate of photosynthesis: • When photosynthesis is high (in light conditions), [DHAP] and [3PG] increase, which inhibits PFK2, which decreases F26BP, which causes a differential increase in F1,6BPase activity over PP-PFK-1), which increases F6P for sucrose synthesis as well as Pi for the continuation of the light reactions. This allows sucrose synthesis when excess DHAP and 3PG occur in the light reactions, which makes great biological sense. • When photosynthesis is low (in dark conditions), the same regulations lead to an increase in F2,6BP, which leads to the preferential activation of the glycolytic enzyme PPi-dependent phosphofructokinase-1 (PP-PFK1) and inhibition of the gluconeogenic enzyme fructose 1,6- bisphosphatase (FBPase-1) We will see later, the main regulatory step in starch synthesis is ADP-glucose pyrophosphorylase. In contrast to the inhibition by 3PG of PFK2, 3PG (which increases in active photosynthesis) activates ADP-glucose pyrophosphorylase while Pi inhibits it. Pi increases when the synthesis of ATP from ADP and Pi (by ATP synthase in the light reaction) slows (such as in darker conditions). If sucrose synthesis slows and sufficient 3PG persists, the activation of ADP-glucose pyrophosphorylase stimulates starch synthesis. Glycosyltransferases (GTs) Glycosyltransferases are very important enzymes as they are involved in the synthesis of most of the biomass on the planet. They catalyze the transfer of a monosaccharide from a donor that has been activated by the attachment of a nucleotide in the form of a nucleotide sugar (NDP-sugar) or dolichol-(pyro)phosphate sugar to acceptors. These include other sugars, lipids, and even proteins, which get glycosylated on alcoholic side chains (Ser, Thr) or amides (Asn). There are over 500,000 different GTs known and deposited in the Carbohydrate-Active enZYmes Database (CAZy database2). Based on sequence and structure there are over 114 different families. Although they depart significantly in primary sequence, only 3 major folds are predominant (GT-A, -B, and -C) a glycosyl transferase reaction is required for the transfer of glucose from a donor like UDP-glucose to an acceptor like fructose to form sucrose as shown in the reaction below. NDP-Glc (donor) + F6P (acceptor) → Sucrose-6-P + NDP Perhaps now is a good time to study their generic reaction mechanisms before we move on to starch synthesis. About 65% of glycosyltransferase reactions use nucleotide sugars as donors. These enzymes are called Leloir transferases. They are nucleotide-dependent. The activated NDP-sugar donor binds first to the enzyme, followed by the acceptor, to form a ternary complex. A conformational change allows catalysis to occur, which leads to the sequential release of products. The enzyme hence follows a sequential ordered bi-bi mechanism. The reaction could proceed with either retention or inversion of the anomeric carbon of the donor NDPsugar. This is illustrated for the reaction of a C1 α-NDP donor monosaccharide with a monosaccharide acceptor to produce the α(1,4) link with retention of configuration or the β(1,4) link with inversion as shown in Figure \(5\). Figure \(5\): Reaction of a donor NDP-monosaccharide and an acceptor monosaccharide with retention or inversion of configuration at the anomeric carbon of the donor The same stereochemical outcomes can occur in the hydrolysis of acetal bonds by glycosyl hydrolases. Reactions that proceed with inversion react in an SN2 reaction, similar to the nucleophilic attack on alkyl halides. For the glycosyl transferase that proceeds with inversion, the attacking nucleophile on the acceptor is made more nucleophilic by general base catalysis by a deprotonated glutamic or aspartic acid. The glycosyl transferase that proceeds with the retention of configuration is less understood. Several alternative mechanisms have been proposed for both inverting and retaining glycosyltransferase in general, as illustrated in Figure \(6\). They include the following possible mechanisms: Panel (A): A double displacement mechanism utilizing two inversions with net retention of stereochemistry involving a covalent glycosyl-enzyme intermediate. The individual steps are inverting via (B) an SN2 process. Panel (B): Inverting Leloir glycosyltransferases promote a backside nucleophilic attack on C1 by the acceptor from an inline (usually equatorial) position, resulting in inversion of the anomeric bond stereochemistry. Panel (C): An orthogonal mechanism consisting of nucleophilic attack on C1 by the acceptor concurrent with leaving group loss from a position approximately at right angles to the C1-leaving group axis. Panel (D): An SNi mechanism involving an intermediate with oxocarbenium character followed by rapid internal nucleophilic attach by the acceptor nucleophile; or Panel (E): An SN1 mechanism involving a discrete oxocarbenium intermediate. All mechanisms require proton transfers of the hydroxyl hydrogen of the acceptor to an enzymatic base or the departing leaving group You will remember from your studies of chemistry that SN1 reactions are dissociative and form a positively charged carbocation intermediate. In a SNi, reaction, the intermediate has cation character but the intermediate is not fully charged. In the case of glycosyltransferase, the intermediates would be oxycarbeniums. (Carbenium ion can be considered carbocations with 3 bonds to the carbon). SN1 reaction will occur only if the formation of the ion is activated and they are stabilized. A protic solvent is typically required for stabilization if the reaction occurs in solution. In the anhydrous active site of the enzyme, an appropriate arrangement of backbone and side chain negative or partially negative atoms is required to provide stabilization for SN1 and SNi mechanisms. The double replacement reactions (Panel A) require a side chain nucleophile and likely candidates in retaining glycosyltransferase are not positioned for such a task. The evidence seems to support the orthogonal mechanism. It appears that the binding of the donor is similar in retaining transferase such that it is in a 900 position of nucleophilic attack by the acceptor, which leads o a trigonal bipyramidal transition state with the nucleophile axial the leaving group equatorial (orthogonal). The donor NDP-monosaccharides typically are Mn2-containing proteins with the inverting and retaining transferase having different coordination geometries for Mn2+ binding, as illustrated in Figure \(7\). Figure \(7\): Coordination geometries for Mn2+ binding in glycosyltransferase. Shuman, ibid. Inverting enzymes such as GalT1 (top) achieve nearly perfect octahedral geometry about the coordinated metal ion (displayed angles of 81° and 91° compared to ideal octahedral 90° bond angles) with subsequent “inline” (approaching 180°) placement of the acceptor nucleophile for classic inverting SN2 backside attack. Retaining enzymes such as GTA (bottom), however, use an arrangement of acidic residues, often with acute bidentate Asp coordination, which severely skews metal geometry (displayed angles of 54° and 115°) and allows sufficient room between phosphate oxygens for orthogonal attack from the acceptor. Uis uridine, C1is donor galactose C1. Figure \(8\) shows a possible mechanism for the transfer of a monosaccharide from the donor ADP-sugar through an oxycarbenium intermediate to an acceptor. Figure \(8\): Possible mechanism for the transfer of a monosaccharide from the donor ADP-sugar through an oxycarbenium intermediate to an acceptor (example - a growing starch chain). Schuman B, Evans SV, Fyles TM (2013) Geometric Attributes of Retaining Glycosyltransferase Enzymes Favor an Orthogonal Mechanism. PLoS ONE 8(8): e71077. doi:10.1371/journal.pone.0071077. Creative Commons Attribution License As mentioned above, there are three major folds for glycosyltransferases, GT-A, GT-B, and GT-C. Different representations of the structure of the GT-A fold core predicted through analysis by neural networks and deep learning are shown in Figure \(9\). Figure \(9\): Fold core of GT-As. Taujale, R., Zhou, Z., Yeung, W. et al. Mapping the glycosyltransferase fold landscape using interpretable deep learning. Nat Commun 12, 5656 (2021). https://doi.org/10.1038/s41467-021-25975-9. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/. This computational modeling of structure uses simple secondary structure representations generated from primary sequences to predict folds irrespective of sequence. GT folds are predicted with high accuracy by learning secondary structure features free of primary sequence alignments. Panel (a) shows a linear map of the conserved secondary core structures (below) and graphs (top). The blue line represents a conservation score and the red a CAM score. CAM values correspond to residue positions that distinguish them the most from other class labels (folds and families). Panel (b) shows a structural alignment of the core with the CAM values mapped onto it. The conserved regions are shown to have a high CAM value indicated by a high intensity of green and a low CAM value of purple. Panel (c) shows on the left the consensus secondary structure for the aligned positions in the two GT-A fold clusters (blue: beta-sheets; red: helices; green: loops). Average CAM values from using different "layers" of analysis are shown for each aligned position (higher intensity of green corresponds to a higher CAM value). Cyan and magenta boxes highlight the secondary structure differences between the two clusters near the hypervariable HV2 and HV3 region respectively. The conserved DXD motif, G-loop, and C-His are indicated for reference. Donor and acceptor substrates for GT-A0 are shown as sticks This figure is shown to give readers a sense of the complexity of the analysis required to understand and differentiate structure/function features for these structurally similar but complex enzymes. Structure and Enzymatic Activity of Sucrose Synthase (SuSy) These enzymes are usually homotetramers with a monomeric molecular weight of around 90,000. The monomers typically have an N-terminal domain that directs the targeting of the enzyme to a specific location and a C-terminal GT-B domain. It can be regulated by phosphorylation at a serine near 12 in the N-terminal domain, which presumably regulates its cellular location, and other near 170 that affects its degradation. Two glutamates in the C-terminal GT-B domain (E678 and E686) and phenylalanine (680) are essential for the catalytic activity. Sucrose synthase is reversible as is the synthesis of sucrose-6-P from F6P and UDP-glucose can be reversed in the presence of UDP. The enzyme can also use ADP-glucose as a donor. The structure of the Thermosynechococcus elongatus sucrose phosphate synthase with bound UDP and sucrose-6-Phosphate has been solved and along with other studies a reaction mechanism proposed as shown in Figure \(10\). Figure \(10\): Catalytic model of TeSPS. Yuying et al. Co-crystal Structure of Thermosynechococcus elongatus Sucrose Phosphate Synthase With UDP and Sucrose-6-Phosphate Provides Insight Into Its Mechanism of Action Involving an Oxocarbenium Ion and the Glycosidic Bond. Frontiers in Microbiology, 11, 2020. https://www.frontiersin.org/article/...icb.2020.01050. Creative Commons Attribution License (CC BY). (A)The state before the reaction is shown. (B)The glucose residue of UDPG forms hydrogen bonds between/among the phosphate groups, His158, Glu331, and F6P. Due to the formation of these hydrogen bonds, the pyranose ring of the glucose becomes negatively charged to promote C1 to form an oxocarbenium ion. (C)The relatively weak hydrogen bond formed by His158 and O6 is broken, which causes the pyranose ring to lose some negative charge character and forces the C1 oxocarbenium ion to form a covalent bond with the F6P oxygen atom. (D)UDP and S6P are released from the catalytic center. A 2D view of the active site residues is shown in Figure \(11\). The catalytic center of TeSPS. Loops 1, 3, 4, and 5 forms a cave that binds to the uracil moiety of UDP. Glu339 stabilizes the ribose ring via the formation of two hydrogen bonds. Leu335 forces two phosphate groups in UDP to reorient. Several basic amino acids, including Arg105, Arg178, Arg249, and Arg253, interact with the phosphate groups of UDP and S6P via ionic bonds. Pro332 at the turn of loop 6 interacts with the pyranose ring via CH/π bonds. All hydroxyl groups (O2, O3, O4, and O6) of the glucose moiety of S6P form hydrogen bonds with phosphate groups or the side chains of various amino acids. “O2” forms a hydrogen bond with P1O1 of the P1 phosphate group of UDP. “O3” forms a hydrogen bond with the carboxyl group of Glu331. “O4” forms a hydrogen bond with P2O1 of the P2 phosphate group of UDP. “O6” forms a hydrogen bond with the imidazole side chain of His158. The distances between groups are indicated in the figure Figure \(12\) shows an interactive iCn3D model of the Thermosynechococcus elongatus Sucrose Phosphate Synthase With UDP and Sucrose-6-Phosphate (6KIH) Figure \(12\): Thermosynechococcus elongatus Sucrose Phosphate Synthase With UDP and Sucrose-6-Phosphate (6KIH) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...1kCnQ9pxcJjdy7 Metabolism of Sucrose Sucrose is a primary synthesis product of photosynthesis and is transported to other plant "sink" tissues where it is used for both energy and biosynthetic precursors. Suc transporters can move it from the phloem to the apoplast. It can enter sink cells through Suc transporters or be hydrolyzed by cell-wall invertase (cwINV) to yield glucose (Glc) and fructose (Fru), which enter by hexose transporters. Suc can also pass directly from the phloem to sink cells through plasmodesmata (physical connections between cells). Inside sink cells, Suc can be metabolized or transported to the vacuole, where it can be stored as Suc, transformed into fructans by fructosyltransferases (FTs), or hydrolyzed by vacuolar invertase (vINV) and stored as hexoses. To be metabolized, Suc must be hydrolyzed by either cytosolic invertase (INV) into glucose and fructose, or by the reversible reaction of sucrose synthase (SuSy) using UDP instead of water to yield fructose and UDP-G. These processes are illustrated in Figure \(13\). The hexoses (glucose and fructose) can be phosphorylated to hexose phosphates (hex-P), directed to starch synthesis in the plastid or glycolysis, and then respiration in the mitochondria or directed to other metabolic pathways. Plasma membrane-associated SuSy (pmSuSy) and cwSUS can generate UDP-G that is used in the synthesis of cellulose for cell walls and callose for plugging plasmodesmata. Callose is a polysaccharide in the form of β-1,3-glucan with some β-1,6-branches in the cell walls of a wide variety of higher plants Biosynthesis of Starch During active photosynthesis in bright light, a plant leaf produces more carbohydrates (as triose phosphates) than it needs for generating energy or synthesizing precursors. The excess is converted to sucrose and transported to other parts of the plant, to be used as fuel or stored. In most plants, starch is the main storage form, but in a few plants, such as sugar beet and sugarcane, sucrose is the primary storage form. The synthesis of sucrose and starch occurs in different cellular compartments (cytosol and plastids, respectively), and these processes are coordinated by a variety of regulatory mechanisms that respond to changes in light level and photosynthetic rate. Starch Synthesis Starch, like glycogen, is a homopolymer of D-glucose in (α1,4) linkage with (α1,6) branches. Glycogen is found in Archaea, Bacteria, and Eukaryotes. In contrast, starch is found in photosynthetic algae, land plants, and in some cyanobacterial species. Starch is synthesized by starch synthase in chloroplasts for temporary storage and in amyloplasts in seeds, roots, and underground stems (tubers) for long-term storage. As with the synthesis of glycogen, the glycosyltransferase catalyzes the addition of an activated ADP-glucose to the acceptor, the elongating starch polymer. ADP-α-D-glucose-1-phosphate (donor) + [(1→4)-α-D-glucosyl](n) (acceptor) ↔ [(1→4)-α-D-glucosyl](n+1) + ADP + H+ The ADP-glucose donor is formed in the following reaction: Glucose 1-phosphate + ATP ↔ ADP-glucose + PPi In plastids, there is, in contrast to the cytosol, a pyrophosphatase which makes the reaction irreversible. Hence the next overall reaction is Starchn + glucose 1-phosphate + ATP → starchn + 1 + ADP + 2Pi Taking into account the hydrolysis by inorganic pyrophosphatase of the PPi produced during ADP-glucose synthesis, the overall reaction for starch formation from glucose 1-phosphate is: Starchn + glucose 1-phosphate + ATP → starchn + 1 + ADP + 2Pi; ΔG'º = -50 kJ/mol = -12 kcal/mole (-50 kJ/mol) ΔG'º = -50 kJ/mol = -12 kcal/mol In glycogen synthesis, the donor is UDP-glucose, and it is added to the reducing end (C1) of the growing glycogen polymer (with the C1 OH acting as a nucleophile) so the polymer extends from that end. Kinetic models suggest that starch synthases, which use ADP-glucose, may use two different active sites that appear to alternately add glucose to the nonreducing C4 end (with the C4 OH acting as a nucleophile), with the reducing end of the linear α(1,4) polymer being alternately covalently attached to one site, then the other, with the attachment activating that end for reaction with C4-OH of the polymer at the other site. Figure \(14\) shows a very simplified structure of the starch synthase using two different active sites as the reaction proceeds through the first steps. Figure \(14\) shows a simplified structure of the starch synthase using two active sites Note that additional glucose units are added to the nonreducing C4 end denoted by a star. α(1,6) branches are added by branching enzymes as in the case of glycogen. Bacteria starch is made in a fashion similar to glycogen, but they use ADP-glucose as do plant starch synthase. Structural models show that the protein has one active site so the kinetic models suggesting the use of two active sites may refer to a movement of chains between different monomers in oligomeric forms of the protein. Both the donor (ADP-Glc) and product (starchn+1) have their participating electrophile (ADP) and now substituted nucleophile (C4-O-R) in the α-anomeric form so they are retaining glycosyltransferases. The enzymes could act in two different ways: • processive mechanism: the acceptor (starchn) stays bound to the enzyme after each addition of the next glucose from the donor ADP-glc, and one chain extends quickly • distributive mechanism: the enzyme dissociates from the product (starchn+1) after the addition of the glucose from ADP-glc, and must rebind to catalyze the next addition, so many new chains start and the growth of each chain is slow Kinetic evidence suggests that some starch synthases are processive and others are distributive. Along with starch synthase, three other enzymes are involved, ADP-glucose pyrophosphorylase (AGPase), starch branching enzyme (SBE), and starch debranching enzyme (DBE) Starch synthesis is regulated by gene transcription, phosphorylation, and redox conditions. A key regulatory enzyme is an ADP-glucose pyrophosphorylase (AGPase). This enzyme catalyzes the formation of ADP-glucose (the donor) and PPi from glucose 1-phosphate and ATP. Dithiothreitol, a reducing agent, increases starch synthesis by inactivating AGPase. We will see below that a key disulfide bond is some starch synthases that must be reduced (cleaved) to open an active site cleft between the N-terminal and C-terminal lobes of the catalytic domain. In vivo, thioredoxins are probably involved in redox regulation. Structures of starch synthetases (SS) The are four classes of soluble starch synthetases (SSI-SSIV) and one starch granule-bound one (GBSS). All have two catalytic domains as noted in the mechanism above except SSII which has 3 CHO binding domains. GBSS appears to form amylose and long chains of amylopectins (amylose with around 5% α(1,6) branching. Loss of function mutants of GBSS have much-reduced amylose concentrations. SSI-SSIII produces most of the amylopectin. SSI is most active with stands (outer ones in branched structures) that have around 7-9 glucoses. In leaves, it generates chains of up to about 10 residues in amylopectin. The SSs can be chloroplastic or amyloplastic. Figure \(15\) shows an interactive iCn3D model of barley starch synthase I in complex with maltooligosaccharide (4HLN). Figure \(15\): Barley starch synthase I in complex with maltooligosaccharide (4HLN). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...LGwFUeJAMskBcA The protein is shown in gray in an open conformation a glucose 5-mer bound on the outside. The protein has a GT-B fold with distinct N- and C-terminal Rossmann-like domains and a central linker. Side chains within 5 Å of oligosaccharide are shown in sticks and labeled. Note especially the disulfide bridge between cysteines 126 and 506 in the central part of the protein that prevents the formation of the active site. This clearly shows the importance of redox signaling to activate the enzyme. The maltose is not bound in the active site but at a surface secondary binding site (SBS). The role of this site is a bit unclear but may be involved in carbohydrate:carbohydrate interactions. Specifically, they may assist in recruiting starch chains for further elongation. SBSs are found in many but not all starch synthases. Note that in this structure there is only one occluded active site and not two as suggested in Figure 14 above. Figure \(16\) shows an interactive iCn3D model of the catalytic domain of starch synthase IV from Arabidopsis thaliana bound to ADP and acarbose (6GNE). Figure \(16\): Catalytic domain of starch synthase IV from Arabidopsis thaliana bound to ADP and acarbose (6GNE) https://structure.ncbi.nlm.nih.gov/i...MioUaKM73M3Wd9 This structure is just the catalytic domain (representing about half of the total protein sequence). The N-terminal domain is colored by the secondary structure. Acarbose (spacefill, CPK colors) again occupies both the donor and acceptor sites in the active site (central regions). The structure has a secondary binding site (SBS) occupied by the disaccharide maltose. Again there are not two active sites, but the migration of starch chains between monomers in oligomeric forms could support the model shown in Figure 14. The structure of beta-acarbose, an inhibitor, is shown in Figure \(17\). Figure \(\PageIndex{x18}\) below shows an interactive iCn3D model of the Granule Bound Starch Synthase from Cyanobacterium sp. CLg1 bound to acarbose and ADP (6GNF) Figure \(18\): Granule Bound Starch Synthase from Cyanobacterium sp. CLg1 bound to beta-acarbose and ADP (6GNF). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?F1tJBDAh6QRhxpH88 ADP, occupying part of the donor site that would usually bind NTP-Glc, is shown in CPK-colored sticks. The rest of the donor site (where Glc of NTP-Glc would bind) and the acceptor site (growing starch chain) is occupied by beta-acarbose, It is shown in the CPK-colored wire-frame surface. Histidine 181, which probably stabilizes an " oxocarbenium-like anomeric carbon in the transition state", is shown in ball and sticks with CPK colors. Cellulose Synthesis We can't leave plant carbohydrate metabolism without considering the synthesis of cellulose, which is catalyzed in plants by members of the superfamily cellulose synthase (CesA) and cellulose synthase-like (CsI) enzymes, which are part of the glycosyltransferase GT2 family and have similar structures. Cellulose and hemicellulose are, of course, chief components of the 10 and 20 cell walls. Members of the CesA family have a conserved motif (DDDQxxRW) as well as a zinc-finger domain. Different members catalyze the synthesis of the 10 and 20 cell walls. Members of the Csl family are involved in additional cell wall glycans including (1,4)-β-D-mannan (CsIA) and xyloglucan cytoskeleton (CslC). UDP-glucose is the donor in the creation of the β(1,4) acetal linkages between glucose monomers. Plant growth must respond to environmental triggers through a balance of cell expansion and cell division and a key regulator of these processes is the flexibility of the cell walls which can maintain turgor pressure by expansion. Cells that are nonexpanding (for example those that line the xylem vessels and in woody tissue) have secondary cell walls beneath their primary walls. We have previously discussed the structure of the primary and secondary cell walls in Chapter 7.3. In brief, the primary cell walls contain cellulose, hemicellulose, and pectins. Cellulose, the main component that provides strength, is synthesized by CesA which forms a very large rosette-shaped complex (CSC). These complexes seem to move intracellular microtubules which guide that synthase complex through the interactions of microtubule-associated cellulose synthase compartments (MASCs), whose numbers increase during stress. Likewise, there are uncoupling proteins that inhibit microtubule movement from the CSC. These protein complexes stay aligned during cell growth. The cell wall hence is a key player in signal transduction that allows growth and cell division. The structure of the rosette-shaped complex (CSC) has been determined by cyroEM and is shown in Figure \(19\). Figure \(19\): Structural cartoons of the CESA CSC complex. Nixon, B., Mansouri, K., Singh, A., et al. Comparative Structural and Computational Analysis Supports Eighteen Cellulose Synthases in the Plant Cellulose Synthesis Complex. Sci Rep 6, 28696 (2016). https://doi.org/10.1038/srep28696. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ Panel (A) shows the complex as a series of trimers of CESAs, with each monomer spanning the membrane with 7 alpha helices. The catalytic domain is in the cytoplasm. Panel (B) shows the CSC complex with the top leaflet removed Panel (C) shows a top-down view showing 6 sets of trimers of CESA. Panel (D) shows a metal replica viewed in the TEM after the removal of the biological material. Each trimer synthesizes a cellulose strand. There are 18 trimers in the complex allowing the concomitant synthesis of cellulose strands that can easily self-associate through hydrogen bonding to form near the extracellular surface cellulose protofibrils.Figure \(20\) shows an interactive iCn3D model of the catalytically active homotrimeric poplar cellulose synthase (6WLB) Figure \(20\): Catalytically active homotrimeric poplar cellulose synthase (6WLB). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...zCm6fYSwAFA897 Each of the monomers within the trimers is given a different color. A 5 residue β(1,4) glycan is shown in cartoon form emerging into the middle of the membrane complex. Figure \(21\) shows an interactive iCn3D model of the homotrimeric poplar cellulose synthase isoform glycan binding site (6WLB). Figure \(21\): Homotrimeric poplar cellulose synthase isoform glycan binding site (6WLB. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...HqYhTQfuw857PA The actual transmembrane channels start just above the active site. Key amino acid side chains (Trp 718, Phe 513, Val 529, and Gln 494) help form the portal opening. The actual channel is lined with both aromatic and hydrophilic residues, which supply sufficient but not too strong noncovalent interactions that allow sequential movement of the continually-synthesized cellulose as it ratchets forward toward the extracellular side of the membrane. The aromatic residues interact through pi stacking with the glucose residues and through interactions with equatorial OH groups on the β-glucose polymer. Remember that cellulose is especially stable from a steric perspective since all its OH groups and the acetal linkage are equatorial.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/20%3A_Photosynthesis_and_Carbohydrate_Synthesis_in_Plants/20.06%3A_Biosynthesis_of_Starch_Sucrose_and_Cellulose.txt

Search Fundamentals of Biochemistry Introduction We present the full fatty acid synthase pathway based on the structure of the yeast fatty acid synthase (FAS) complex, whose full structure is known. Then we will explore each step in more detail. The mammalian FAS complex is a bit different and we will summarize those differences next. We will end with a summary of the bacterial pathway. Let's look at the net reaction first and then go back to the structure and mechanism from that. Equation 1 below shows the net reaction. $8 \text { Acetyl-CoA }(2 \mathrm{C})+14 \mathrm{NADPH}+14 \mathrm{H}^{+}+7 \mathrm{ATP} \rightarrow 1 \text { Palmitate }(16: 0)+8 \mathrm{CoA}+6 \mathrm{H}_{2} \mathrm{O}+14 \mathrm{NADP}^{+}+7 \mathrm{ADP}+7 \mathrm{Pi}$ Eight 2C-acetyl-CoAs condense to make the 16:0 fatty acid palmitate. That acetyl-CoA is used to synthesize and elongate fatty acids suggests an immediate explanation for the fact that most fatty acids have an even number of carbon atoms. The net reaction is a reductive biosynthesis making one long molecule from multiple short ones. Hence there must be an energy source to drive the reaction (7ATPs) and a reducing agent (typically NADPH). This reaction takes place in the cytosol and uses the nicotinamide-based redox pair (NADPH/NADP+). These factors differentiate fatty acid synthesis from beta-oxidation, which occurs in the mitochondria and which uses the redox pair NAD+/NADH. From a chemical perspective, the acetyl-CoAs, high-energy molecules with respect to their hydrolysis products, must be activated using ATP in some way to drive C-C bond formations. That reaction is catalyzed by the enzyme acetyl-CoA carboxylase, which uses the ATP already mentioned and the bound cofactor biotin that adds CO2 to acetyl-CoA to form malonyl-CoA. Neither malonyl-CoA nor CO2 is shown in the above equation since they are present on both sides of the net reaction and cancel out from the final balanced equation. C-C bond formation occurs on the addition of the growing acyl group with the 3C malonyl with the actual addition reaction driven by the release of CO2. The malonyl-CoA produced by acetyl-CoA carboxylase feeds now into the fatty acid synthesis cycle. Only one other enzyme complex is required for the entire reaction, the fatty acid synthase (FAS) complex. Don't let that fool you into thinking the mechanism is simple though! There are two types of fatty acid synthase complexes, I and II. Type II FAS is found in plants, most bacteria, and in mitochondria. (Its presence in the mitochondria might seem odd since fatty acid oxidation occurs there. We will discuss that in a bit.) There are multiple separate enzymes to catalyze the synthesis of fatty acids in the Type II FAS systems. We won't discuss those. Instead, we will focus on the Type I FAS complex found in some bacteria, fungi like yeast, and higher eukaryotes. The Type I FAS is one very large complex. In mammals, it consists of two α chains with a total molecular weight of 540,000. In yeast, it is an α6β6 heterododecamer with a molecular weight of 2.6 million. In either complex, the alpha and beta chains have multiple different enzyme catalytic domains. Large enzymatic complexes typically allow bound substrates to proceed to products without dissociation, with the bound intermediates moving through "channels" to the next active site. In the case of Type I FAS, the growing intermediate, which eventually reaches the end product size 16:0 is tethered to an acyl-carrier protein (ACP). The tethered intermediate can "swing" from one active site to another to allow the iterative stepwise chemical reactions to complete before the addition of another 2C acetyl CA to the growing chain. The actual extra 2Cs come from the 3C malonyl-CoA (formed by the extracyclic (outside of the pathway) enzyme acetyl-CoA carboxylase) and immediate release of CO2 which helps drive the reaction forward. Now we can introduce the fatty acid synthase cycle and feeder acetyl-CoA carboxylase reaction in its entirely, as shown in Figure $1$ for yeast FAS1. This is a difficult figure to understand but keep in mind that it scales down the complexity of FAS1 enormously. Let's deconstruct the figure to make it more understandable. The dotted red boxes show the substrates and products that match chemical equation 1. The substrate acetyl-CoA comes into the cycle in two places shown in the extracyclic section to the upper left. It comes in as acetyl-CoA from reaction 3 after attachment to the acyl carrier protein (part of FAS1). It also comes in "effectively" after it has been carboxylated by acetyl-CoA carboxylase in reaction 2 to malonyl-CoA, which enters the cycle as malonyl-ACP after reaction 5. The other substrates/products include CoAs the NAD(P)H/NADP+ couples and H2O shown around the right-hand side of the circular fatty acid synthesis cycle (2 -6 o'clock positions) as well as the final product, 16:0-ACP (9:30 clock position), which forms 16:0-SCoA. The individual enzymatic domains in the α6β6 hetero-dodecamer yeast FAS1 are shown in colored spheres as shown in Figure $2$. Figure $3$ summarizes pictorially the repetitive Claisen condensation reactions of acetyl-ACP and malonyl-ACP in the first step, and the growing acyl-ACP chain and malonyl-CoA in the next two cycles. In each step, the condensation (C-C bond formation) is driven by the decarboxylation of malonyl-CoA. (Of course, earlier expenditure of ATP in step 1 is required). Electrons in malonyl-CoA that form a bond with acetyl-ACP in the first condensation step and acyl-ACP in subsequent condensations are shown as bold blue lines. Note that the condensation is from the head of the malonyl-ACP to the tail of the elongating acyl chain. The ACP or equivalent chains are in the interior of the complex which allows movement of the acyl groups attached to pantetheine chains to interactively reach nearby catalytic domains for each step in the cycle. Mechanism of individual reactions Now we can explore some of the reactions in more detail. Then we will look at the structure of yeast FAS and its domain organization. Reaction 1: Phosphopantetheine transferase Holo-(acyl carrier protein) synthase (AcpS) from Bacillus subtilis is a member of the phosphopantetheinyl transferase superfamily. AcpS post-translationally modifies ACP to its holo form to activate it. AcpS catalyzes the transfer of the 4'-phosphopantetheinyl (P-pant) moiety of coenzyme A to a serine residue on the ACP. This gives the activated ACP enzyme and adenosine 3'5'-bisphosphate as products. This process is important as ACP enzymes play important roles in several biosynthetic pathways, such as the synthesis of fatty acids, and vitamins, AcpS is essential in the initiation of the biosynthesis of fatty acids, polyketide antibiotics, and non-ribosomal peptide Figure $4$: https://www.ebi.ac.uk/thornton-srv/m-csa/entry/152/ Reaction 2: Acetyl-CoA Carboxylase (ACC) This enzyme catalyzes the carboxylation of acetyl-CoA to malonyl-CoA. which then enters the fatty acid synthesis cycle. This is a key enzyme as it is the rate-limiting step, and is regulated. The reaction has two steps: • carboxylation of biotin on a biotin carboxyl carrier domain of the enzyme at the expense of ATP hydrolysis. The carboxylation, using bicarbonate as a substrate, leads to the formation of a molecule with high energy (with respect to its hydrolysis product), the carboxylated biotin intermediate.  (Remember, there is no such thing as a "high energy" bond.) • transfer of the carboxyl group to acetyl-CoA to form malonyl-CoA which requires the formation of a C-C covalent bond. Biotin, an essential nutrient and cofactor, "carries" activated carboxyl groups for transfer. It is a carboxyl (not acetyl) donor. It is linked to a lysine side chain in the protein through an amide link. The enzyme is downregulated by 16:0-SCoA, the end product of the pathway for fatty synthesis by FAS, and by phosphorylation by kinase activation through the cAMP pathways from glucagon binding. You don't want to synthesize fatty acids when your energy state is low (signaled by increases in glucagon). Its activity is upregulated by the binding of citrate to an allosteric site. This makes sense since high citrate, a citric acid cycle intermediate, signifies abundant energy reserves are available so fatty acids would be synthesized for future needs. The dephosphorylated form of the enzyme would signify the need to increase fatty acid synthesis. Figure $5$ shows a few steps in the carboxylation of biotin and the subsequent transfer of the carboxy group to acetyl-CoA, forming malonyl-CoA. BCCP represents the biotin carboxyl carrier domain of acetyl-CoA carboxylase. Biotin carboxylases are found in many enzymes and pathways, not just fatty acid synthesis. Scientists are trying to devise new pathways and enhanced carboxylases to pull CO2 from the air into biosynthetic reactions producing fuels, which could be burned. This would release CO2 back into the atmosphere, in a process that would theoretically but perhaps not practically carbon neutral with respect to greenhouse gas emission. The abbreviated mechanism shown above shows two of the three activities of acetyl-CoA carboxylase. They are biotin carboxylase (BC) and carboxyltransferase (CT) activity. The third is the biotin carboxyl carrier protein (BCCP) which links biotin to it through a lysine side chain. Different enzymes have different acceptors (acetyl-CoA, pyruvate, etc) of the activated carboxy group so the structure of those centers varies significantly. The BC and BCCP components are structurally similar for different carboxylases. A mechanism for the E. Coli biotin carboxylase component of the reaction is shown in Figure $6$. Glu 296 acts as a general acid/base while Arg 338 stabilizes negative charge in intermediates and transition states. Structure of acetyl-CoA carboxylase (ACC) There are two forms of ACC, soluble cytosolic ACC1 and mitochondrial membrane-associated ACC2 (the latter which regulates beta-oxidation of fatty acids. The overall domain structure of yeast acetyl-CoA carboxylase is in Figure $7$. Panel (a) shows the domain organization of ScACC. The domains are labeled and given different colors. The five domains of ACC Central (AC1–AC5) are labeled 1–5. The phosphorylation site in the central region is indicated. The phosphorylation site before the biotin carboxylase (BC) domain core is indicated with the dashed lines, as it is absent in ScACC. Panel (b) shows the structure of the ScACC holoenzyme dimer [7]. One protomer is shown as ribbons, while the other as a surface. The domains in the monomers are colored according to panel (a) and labeled. Ser1157 (red star) is located in a loop missing in the structure (dashed lines), and its distances to the BC and carboxyltransferase (CT) active sites (black asterisks) and the BC dimer interface (black rectangle) in the holoenzyme are indicated. A chain domain structure 5CSKA_Yeast Acety-lCoA Carboxylase is shown in Figure $8$. The colors used to show the domains are matched to Figure $\PageIndex{x}$ above and are as follows: • Red - BC, biotin carboxylase (has enzymatic activity) • Gold - BT, an interaction domain • Blue - BCCP (biotin carboxyl carrier protein (disordered blue spheres) • Cyan - AC (central bridging noncatalytic domain comprising AC1-AC5). • Yellow - CT, carboxyl transferase (has enzymatic activity), comprising the N and C subdomains A similar domain structure is found in human ACC1. The central bridging domain in human ACC1 is called the Central Domain (CD) with four parts, CDN, CDL, CDC1, and CDC2. Regulation of ACC ACC1 activity is regulated by product inhibition, allosteric effector, and phosphorylation by different kinases. The enzyme is inhibited by its product, malonyl-CoA, and also the end product of fatty acid synthesis, 16:0-CoA. The protein is inactive as a monomer and is active as a dimer. Anything that can perturb that equilibrium can affect ACC activity. Phosphorylation generally inhibits the protein by promoting the dissociation of the active dimer. The active dimer, in the presence of the allosteric activator, citrate can aggregate to form fibrils, which are even more active than the dimer. The protein is post-translationally modified by phosphorylation at several sites but a few stand out. AMP-activated protein kinase (AMPK) phosphorylates human ACC at AMP-activated protein kinase at Ser 80 (in the BC domain). In yeast, the kinase SNF1 (equivalent to human AMPK) does not phosphorylate Ser 80 but it phosphorylates a key Ser 1157 in the AC subdomain AC4 in yeast. • In humans, phosphorylation at Ser 80 in the BC domain prevents dimerization of the protein, through the BC domain and hence inhibits activity; • In yeast, phosphorylation st Ser 1157 in AC subdomain AC4 in yeast produces a conformational change in the BC domain region that appears to lead to dimer dissociation. Human ACC is phosphorylated at a multitude of sites with Ser 80 and 1201 (which is phosphorylated both by AMPK and PKA) being the most important in regulation. In addition, it is phosphorylated at Ser 1263 by a cyclin-dependent protein kinase (CDK). When phosphorylated, the pSer 1263 facilitates the binding of BRAC1, the tumor suppressor protein which when mutated can dramatically increase breast cancer. The binding of BRAC1 leads to an inactive fibril form of ACC. Figure $9$ summarizes the regulation of human CoA carboxylase. 5. Malonyl-Palmitoyl Transferase (MPT)/Malonyl-CoA-acyl carrier protein transacylase (MAT) These protein activities are found in the yeast (MPT) and human (MAT) domains. The mechanism is shown in Figure $10$. Two key catalytic residues, Ser 92 and His 201, are involved in an acylation/deacylation of the catalytic serine. The mechanism is shown in Figure $11$. Reaction 9. Enoyl Reductase This is the last reaction in the fatty acid synthase cycle, which of course repeats until a 16:0-SCoA is made. The yeast enzyme is different in that it has a tightly-bound FMN which is involved in electron transfer (reduction) along with NADPH. The mechanism is likely ping-pong with NADP+ released before the enoyl acyl-CoA binds. Figure $12$ shows an interactive iCn3D model of the enoyl-acyl carrier protein reductase (ER) in complex with NAD+ and triclosan (1QSG) Figure $12$: enoyl-acyl carrier protein reductase (ER) in complex with NAD+ and triclosan (1QSG) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...Z2ZCQaTYabQWa9 Triclosan is used in many commercial products, even soaps, as it has broad-spectrum antimicrobial activity. In a world with increasing antibiotic resistance, it is unwise to use triclosan in soaps since the evidence shows that the surfactant properties of soaps are sufficient to remove bacteria from the skin. Triclosan is an active site inhibitor of bacterial enoyl-reductases but in humans, it appears to be an allosteric inhibitor, since it is bound at a protein-protein interface and not the active site. Structure of Yeast Fatty Acid Synthase I (FAS1) Now you have enough background to explore the actual structure of fatty acid synthase. Animal FAS1 is an α2 homodimer, some bacterial FASs are α6 hexamers, while fungal FASs are α6β6 dodecamers. In both cases, the monomers are multi-domain proteins with each domain having a different catalytic activity. There is another type of fatty acid synthase II (FAS2), that consists of separate enzymes each with their own catalytic function. Type II FAS around found in plants, most bacteria, and in mitochondria. We'll explore the "more interesting" Type I and start with yeast (Saccharomyces cerevisiae) FAS (6QL5), whose structure has been solved with a regulatory γ subunit that 2.8 angstrom 6QL5. Figure $13$ shows an interactive iCn3D model of the fatty acid synthase complex with bound gamma subunit from Saccharomyces cerevisiae (6QL5) very long load time. Figure $13$: Fatty acid synthase complex with bound gamma subunit from Saccharomyces cerevisiae 6QL5. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...YnMNSaTbTXYGe7 (very long load time). The alpha subunits are shown in cyan and the beta subunits are shown in magenta. The regulatory γ subunit is shown in orange. The 6 alpha subunits are arranged in an equatorial wheel or disk. The beta subunits are arranged in trimer top and bottom "domes" that cover the alpha subunit disk. Inside are 3 spaces or chambers where the reactions occur. 5 openings allow outside substrate access and six openings allow internal metabolite access. The FAS is more porous than in large structures with encapsulated volumes like the proteasome or viral capsids. It is more similar to pyruvate dehydrogenase. The ACP protein domain is found inside the structure and is tethered and "swingable" to allow the transfer internal transfer of acyl intermediates to the different enzymatic functional domains of the complex. Figure $14$ shows animated images of yeast fatty acid synthase complex, which should give you a sense of the arrangement of the equatorial alpha-helical wheel, the two beta subunit domes, and the interior and interior volumes available for reaction chemistry and intermediate movement. They switch between the full structure, followed by separate views of the alpha (cyan), beta (magenta), and regulatory (orange) chains. fatty acid synthase complex - side view fatty acid synthase complex - top view Figure $14$: Alpha chains (cyan), beta chains (magenta), regulatory (orange) The α and β monomers are multi-domain proteins with different domains having different catalytic properties. Just color coding the monomers in just one color doesn't give insight into the amazing catalytic properties of each chain. Figure $15$ shows colored-coded domain and protein structures for yeast α6β6 dodecamer. The color-coding for the domains for each subunit shown at the bottom of panel A is also used for the domain structures in the actual protein structures. Panel (A) shows the structure of S. cerevisiae FAS (3hmj). The side (left) and top view (right) with two β-subunits and two α-subunits are shown. Note that ACP is in the FAS interior, but is not shown. The MPT fold is comprised of both subunits and shown in cartoon representation (β part in brown and its α part in red). Nomenclature: acetyltransferase (AT), enoyl reductase (ER), dehydratase (DH), malonyl-palmitoyl-transferase (MPT), an acyl carrier protein (ACP), ketoacyl reductase (KR), ketoacyl synthase (KS) and phosphopantetheine transferase domain (PPT). Insertion elements are highlighted in grey; trimerization module (TM), 6-stranded β-sheet (SBS), hotdog-domain 2 (HDD2), dimerization module 1–4 (DM1-4), 4-helical bundle (4HB)). Please note that DM2 is not visible in this structure. Panel(B) shows three yeast FAS barrels as a central D3-symmetric α hexamer (α6-wheel) and two C3-symmetric β trimers (β3-domes). β3-domes have been shifted for clarity (see arrows). ACP domains are shown for two α-subunits and are modeled by spheres in magenta. ACP linkers are indicated by dashed lines. Panel (C) shows the post-translational modification of ACP. For phosphopantetheinylation, ACP and PPT have to physically interact. Fischer, M., Joppe, M., Mulinacci, B. et al. Analysis of the co-translational assembly of the fungal fatty acid synthase (FAS). Sci Rep 10, 895 (2020). https://doi.org/10.1038/s41598-020-57418-8. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ In Figure $16$ we connect the domain structures of each monomer to the actual catalytic cycle of yeast FAS. Figure $16$: Domain structure and the catalytic cycle of yeast fatty acid synthase (adapted from Singh et al. Cell, 180 (2020), https://doi.org/10.1016/j.cell.2020.02.034 and Fischer et al. Sci Rep 10, 895 (2020). https://doi.org/10.1038/s41598-020-57418-8. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ Figure $17$ shows the actual structure of the isolated α and β monomers. The color coding of the domains is the same as used above. Fatty acid synthase 1 (FAS1) beta subunit (FAS1) (6U5U G chain) Fatty acid synthase 2 (FAS2) alpha subunit (2UV8A - P19097 AlphaFold) Figure $17$ shows the approximate domain structures and locations for the yeast fatty acid synthase 1 (FAS1) beta subunit (6U5U G chain) and fatty acid synthase 2 (FAS2) alpha subunit. The color coding for the domains in each subunit is as follows: FAS1 beta subunit: AT, ER, DH, MPT, flavin mononucleotide (FMN) spacefill cyan, NADP nicotinamide-adenine dinucleotide phosphate (NAP), spacefill magenta;  FAS2 alpha subunit: ACP, KR, KS, PPT Now, let's zoom in to see the actual interactions of bound 4'-phosphopantetheine and flavin mononucleotide (FMN). Phosphopantetheine is bound between the alpha (cyan) and beta (magenta) subunits as shown in Figure $18$. 4'-phosphopantetheine between alpha (cyan) and beta (magenta) subunits flavin mononucleotide (FMN) within beta (FAS1) subunit Figure $18$: The γ subunit crosses the entire inner cavity of FAS interfering with the activities of the reductases. A central difference in the FAS1 found in humans and other animals occurs in the first reaction in which the acetyl group of acetyl-CoA is transferred to the pantetheine sulfhydryl of the acyl carrier protein (ACP) domain catalyzed by the malonyl‐/acetyltransferase (MAT) domain. It is then transferred to the active site cysteine of the β‐ketoacyl synthase (KS) domain. This allows the malonyl group to be transferred to the now free ACP domain for a second transfer reaction. A Claisen condensation reaction driven by the decarboxylation of the malonyl-ACP occurs to form the β‐ketoacyl intermediate bond to the KS domain. An alternative view of the mammalian fatty acid cycle is shown in Figure $19$. Panel (a) shows the priming of animal fatty acid synthesis. Panel (b) shows how in the first step, the substrate is selected by the MAT domain and transferred to the ACP domain (Step 2) from where it is passed on to the KS domain (Step 3). Important active site residues are highlighted and C161 is marked with an asterisk. The porcine FAS structure (2png) is shown. Domains of one protomer of FAS homodimer are colored. ACP, acyl carrier protein; DH, dehydratase; ER, enoyl reductase; FAS, fatty acid synthase; KR, ketoreductase; KS, ketosynthase; MAT, malonyl‐/acetyltransferase; PPant arm, expand fully in the figure; TE, thioesterase Figure $20$ shows the yeast FAS cycle (as a proxy for the mammalian cycle) placed in context with the mitochondrial matrix pathways that feed acetyl-Coa and NADPH into the fatty acid synthesis pathway. Figure $20$: yeast FAS cycle (as a proxy for the mammalian cycle) placed in context with the mitochondrial matrix pathways that acetyl-Coa and NADPH into fatty acid synthesis Fatty acid Elongation and Desaturation Overview Elongation of fatty acids occurs in the cytoplasm of mammals from 16:0 made through fatty acid synthase. As with the synthesis of 16:0 by FAS, the elongation consists of two carbon addition driven by the decarboxylation of a malonyl-CoA substrate. Elongation is carried out by a family of ELOngation of Very Long-chain fatty acid enzymes (ELOVLs). Desaturation is catalyzed by Stearoyl-CoA Desaturases (SCD1 and SCD2) and Fatty Acid Desaturases (FADS1 and FADS2). Figure $21$ shows an overview of de novo fatty acid synthesis with elongation and desaturation in mammals as well as elongation and desaturation of dietary fatty acids. FAs are elongated (ELOVL1–7) and/or desaturated (SCD, FADS) to generate complex FAs. Long-chain saturated FAs (LCSFA) and unsaturated FAs of ω9 and ω7 can be synthesized from palmitic acid (PA, C16:0) produced by the de novo FA synthesis. Long-chain unsaturated FAs of the ω6 and ω3 series can only be synthesized from essential diet-derived FAs (OA, oleic acid; NA, nervonic acid; ALA, α-linolenic acid; DHA, docosahexaenoic acids; LA, linoleic acid; ARA, arachidonic acid). SFA are saturated fatty acids. Elongation Mammals use endoplasmic reticulum very long-chain fatty acid enzymes (ELOVLs). There appear to be seven in mammals with different substrate specificities as shown in the figure above. The nomenclature for fatty acids is a bit strange. Those with 11-20 carbons are called long-chain FAs (LCFAs) and those with more than 20 carbons are very long-chain FAs (VLCFAs). Those with more than 26 carbons are called ultra long-chain FAs (ULCFAs), which are found in the skin, retina, meibomian gland, testis, and brain. The same four steps (condensation, reduction, dehydration, and reduction) used in FAS are used in elongation. The enzymes are abbreviated a bit differently (3-ketoacyl-CoA reductase or KAR, 3-hydroxyacyl-CoA dehydratase or HACD, trans-2-enoyl-CoA reductase or TER and LCFA). The enzyme is separate and not part of a multifunction complex like FAS. The elongation cycle is shown in Figure $22$. Figure $23$ shows fatty acid composition in sphingomyelins which typically have long fatty acids. Mechanism of ELOVL7 ELOVL condensing enzymes have a long hydrophobic tunnel and an active site nucleophilic histidine. More traditional nucleophiles like serine or cysteine were eliminated as potential candidates for the active site nucleophile since none were in proximity. The active site with a key HxxHH motif lies deep in the membrane. Nucleophilic catalysis leads to an N-acyl intermediate and is consistent with the observed ping-pong kinetics. An alternative reaction for a "bisubstrate" reaction is the formation of a ternary complex with both substrates. The narrow size of the binding pockets precludes this mechanism, so one product, after it is formed, must dissociate before the next substrate can bind. ELOVL7 preferentially catalyzes the elongation of C18-CoA presumably arising from the size of the acyl-binding product. A plausible mechanism for human long-chain fatty acid enzyme 7 (ELOVL7) elongation is shown in Figure $24$ and Figure $25$. Figure $24$ shows the beginning acylation step of the reaction. Figure $24$: Acylation step in the human long-chain fatty acid enzyme 7 (ELOVL7) elongation reaction Figure $25$ shows the condensation (with the elimination of CO2), followed by the reduction and dehydration of human ELOVL7. Figure $26$ shows an interactive iCn3D model of the human ELOVL fatty acid elongase 7 (ELOVL7) with bound 3-keto eicosanoyl CoA (6Y7F). Figure $26$: Human ELOVL fatty acid elongase 7 (ELOVL7) with bound 3-keto eicosanoyl CoA (6Y7F). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...eajMeiRgfHd9s9 The bound 3-keto eicosanoyl CoA is shown in spacefill, CPK colors. Key amino acids involved in catalysis are shown in sticks, CPK colors, and labeled. Desaturation The introduction of double bonds requires a different set of enzymes, acyl-coenzyme A (CoA) desaturases. Mammals have desaturases that can produce double bonds at Δ9, Δ6, and Δ5. There are two types, stearoyl-CoA desaturases (SCDs), which introduce a double bond at C9 of saturated fatty acid, and fatty acid desaturases (FADS), which work on unsaturated fatty acids. These are shown in Figure $20$. Plants, but not mammals, have Δ12 and Δ15-desaturases, so they, but not mammals, can synthesize ω6 and ω3 fatty acids. These must be supplied by the diet as essential fatty acids and are precursors for the synthesis of longer fatty acids like arachidonic acid (20:4Δ5,8,11,14), necessary for prostaglandin synthesis (see below) and docosahexaenoic acids (DHA). A plausible mechanism for the stearoyl-CoA desaturase from castor seeds is shown below in Figures 27-30. The mechanisms and explanations are adapted from https://www.ebi.ac.uk/thornton-srv/m-csa/entry/136/. Creative Commons Attribution 4.0 International (CC BY 4.0) License At the start of the reaction, ferredoxin, a small iron-sulfur protein donates a single electron, through Trp62, Asp228, and His146, to one of the Fe(III) centers in the desaturase. Ferredoxin then donates a second single electron, through Trp62, Asp228, and His146, to the second Fe(III) center. Both of these are shown in Figure $27$ Part 2 of the reaction is shown in Figure $28$. Figure $28$: Part 2 of the stearoyl-CoA desaturase The reduced charges on the Fe ions lead to loss of the oxide bridging ion (O2-) and its replacement with dioxygen. Water coordinates to one of the Fe(II)s, which causes it to donate a single electron to the dioxide molecule. This starts the first of two homolytic additions of the dioxygen molecule to both Fe(II) centers and the second Fe(II) center also donates a single electron to the dioxygen bridge. Part 3 of the stearoyl-CoA desaturaseFigure $29$ is shown below. Part 4 of the stearoyl-CoA desaturase is shown in Figure $30$. In this reaction the hydroxide on the first Fe(IV) center deprotonates the stearoyl-[acyl-carrier protein] substrate, initiating the elimination of a hydride ion, which attacks the second Fe(IV) bound hydroxide. The excess electrons are donated singly to both Fe(IV) centers, regenerating the enzyme. Figure $31$ shows an interactive iCn3D model of the Stearoyl-acyl-carrier protein desaturase from castor seeds (1afr) Figure $31$: Stearoyl-acyl-carrier protein desaturase from castor seeds (1afr) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...UonrwyWfcg9wU8 An electron transport chain carries an electron from ferredoxin to one of the iron centers via several residues. A second electron is then carried to the second iron center. Several redox reactions take place involving oxygen, water, and iron centers, which result in the deprotonation of the substrate and the formation of a double bond. There are two possible paths for the electron transport chain - the one described here has been chosen because of its analogy to what has been suggested for E. coli ribonucleotide reductase protein R2. Mitochondrial FA Synthesis Mitochondria also engage in limited fatty acid synthesis. They use separate discrete enzymes (at least 6) as do most bacteria. They don't employ a multienzyme complex as described above for yeast and mammals. Nevertheless, there are commonalities. Synthesis proceeds by the addition of two carbon units to a growing acyl-ACP chain using malonyl-CoA and the release of CO2 to drive the condensation process (in addition of course to ATP cleavage). The cytoplasmic fatty acid synthesis produces 16:0 (palmitate). In contrast, mitochondrial FAS produces two major products, 8:0 (octanoate). This is then converted to the 8C derivative lipoic acid. Their structures are shown in Figure $32$. We have seen lipoic acid before as a cofactor in pyruvate dehydrogenase and α-ketoglutarate dehydrogenase, two enzymes that catalyze the more difficult decarboxylation of α-ketoacids (which don't have a built-in electron sink at a β carbon adjacent to the departing carboxyl group. It is also used in branched-chain amino acid dehydrogenases. The absence of lipoic acid is lethal in mice. Figure $33$ shows the fatty acid synthesis pathway in mitochondria. Figure $33$: mitochondrial fatty acid synthesis pathway and downstream lipoic acid synthesis. Nowinski et al. eLife 2020;9:e58041. DOI: https://doi.org/10.7554/eLife.58041. Creative Commons Attribution License Some of the enzymes in the pathway include the following: Mcat is malonyl-CoA ACP transacylase, and Oxsm is beta-ketoacyl synthase that catalyzes the Claisen condensation of malonyl-ACP with the growing acyl chain. CBR4 is 3-oxoacyl-[acyl-carrier-protein] reductase. HSD17B8 is (3R)-3-hydroxyacyl-CoA dehydrogenase. HTD2 is hydroxyacyl-thioester dehydratase type 2). Mecr is the terminal enoyl-[acyl-carrier-protein] reductase. LIPT2 is lipoyltransferase 2. LIAS is lipoyl synthase. LIPT1 (Lipoyltransferase 1) catalyzes the terminal step in lipoic acid and lipoylated protein synthesis. It now appears that longer fatty acyl chains (up to 14C) are synthesized in mitochondria. Mutations in key enzymes above have been made that don't affect lipoic acid synthesis but impair mitochondrial function. Eicosinoid and prostaglandin synthesis As shown in Fig 20, arachidonic acid, 20:4Δ5,8,11,14, an ω-6 fatty acid (5,8,11,14- eicosatetraenoic acid), is synthesized from linoleic acid, 18:2Δ9,12, another ω-6 polyunsaturated fatty acid, through a series of elongation and desaturation steps. Arachidonic acid is released from membrane phospholipids on activation of phospholipase A2 and converted to a class of molecules called prostaglandins, which have powerful hormone-like effects. The prostaglandins are part of a group of icosanoids derived from cleavage of membrane ω-3 (20:5Δ5,8,11,14,17, eicosapentaenoic acid) and ω-6 C20 fatty acids, which are metabolized to form leukotrienes (LTs), prostaglandins (PGs), prostacyclins (PCs), and thromboxanes (TXAs). These have differing and sometimes opposing physiologically effects in animals. Prostaglandins have been found in almost every tissue in humans and other animals ω-3 and ω-3 and fatty acids The most common polyunsaturated fats (PUFAs) in our diet are the ω-3 and ω-6 classes. Most abundant in the ω-6 class in plant food is linoleic acid (18:2 ω-6, or 18:2Δ9,12), while linolenic acid (18:3 ω-3 or 18:3Δ9,12,15) is the most abundant in the n-3ω class. These fatty acids are essential in that they are biological precursors for other PUFAs. Specifically, • linoleic acid (18:2 ω-6, or 18:2Δ9,12) is a biosynthetic precursor of arachidonic acid (20:4 ω-6 or 20:4Δ5,8,11,14) • linolenic acid (18:3 ω-3, or 18:3Δ9,12,15) is a biosynthetic precursor of eicosapentaenoic acid (EPA, 20:5 ω-3 or 20:5Δ5,8,11,14,17) and to a much smaller extent, docosahexaenoic acid (DHA, 22:6 ω-3 or 22:6Δ4,7,10,13,16,19). These essential precursor fatty acids are substrates for intracellular enzymes such as elongases, desaturases, and beta-oxidation type enzymes in the endoplasmic reticulum and another organelle, the peroxisome (involved in the oxidative metabolism of straight chain and branched fatty acids, peroxide metabolism, and cholesterol/bile salt synthesis). Animals fed diets high in plant 18:2(n-6) fats accumulate 20:4(n-6) fatty acids in their tissues while those fed diets high in plant 18:3(n-3) accumulate 22:6(n-3). Animals fed diets high in fish oils accumulate 20:5 (EPA) and 22:6 (DHA) at the expense of 20:4(n-6). Figure $34$ We will focus our attention on the conversion of arachidonic acid (20:4 ω-6 or 20:4Δ5,8,11,14) to prostaglandin G2 (PGG2) and H2 (PGH2) by the enzyme cyclooxygenase I and II. These enzymes are also called by their more formal names, prostaglandin endoperoxide H synthases (PGHSs) I and II. COX-1 prefers arachidonic acid as a substrate, while COX-2 has a broader substrate specificity and can use eicosapentaenoic acid (EPA, 20:5 ω-3 or 20:5Δ5,8,11,14,17) and even neutral substrate derivatives of arachidonic acid, including 2-arachidonoylglycerol and anandamide, both ligands for cannabinoid receptors. COX 1/2 use two dioxygens and 2 electrons to make PGH2. Two different, separate but connected catalytic sites are used to catalyze the two steps (shown in Figure $34$) 1. Arachidonic acid (20:4Δ5,8,11,14) is converted to PGG2. This step is catalyzed by a bis-oxygenase domain. The word oxygenase is used since both atoms of dioxygen are added to the reactant. Bis indicates that two different molecules of dioxygen are added. Since one of the dioxygens added forms an endoperoxide cyclic bridge in PGG2, this enzymatic domain, and the entire enzyme is usually called cyclooxygenase (COX). The second dioxygen is added as a peroxide (both oxygen atoms of dioxygen added); 2. Prostaglandin G2 (PGG2) is then converted to prostaglandin H2 (PGH2) in which the noncyclic peroxide form in the first step is converted to hydroxide. This is also a reduction step since the oxidation numbers of the oxygen atoms in the peroxide, -1, change to -2 (a gain of electrons) in the hydroxide in PGH2. This reaction is catalyzed by the peroxidase (POX) activity of the enzyme. A cofactor heme in the POX site facilitates catalysis. The COX reaction proceeds through radical intermediates. but it requires the activation of peroxide by the heme in the POX site. We'll show two different mechanisms for both reactions (COX and POX) catalyzed by the enzyme prostaglandin endoperoxide H synthase, which we will also call COX. The first mechanism shown below in Figure $35$ emphasizes the free radical nature of the reactions. It shows just one amino acid (Tyr 385) involved in the reaction. Note that the first step in the reaction shows the added dioxygen as an excited state "singlet" O2, but it is parenthetically noted that it most likely involves O2 binding to and being activated by the heme iron in POX site. A more detailed reaction mechanism for the synthesis of PGG2 by prostaglandin endoperoxide H synthase (COX), emphasizing the role of active site amino acid and the heme in the POX site, is shown in Figures 36-38 below. (The mechanism and its explanation are adapted from https://www.ebi.ac.uk/thornton-srv/m-csa/entry/37/. Creative Commons Attribution 4.0 International (CC BY 4.0) License) The 1st step involves a peroxide (denoted H-O-O*) but it might also be a ROO peroxide or NO. The generation of a Tyr-385. free radical in the COX site. is critical for the reaction. This is preceded by the heme radical cation in the POX site. Figure $36\ below shows the first step in which His 207 deprotonates the alkyl peroxide, which then coordinates to the heme Fe(III) in the POX site. In the next step, the peroxy bond donates two electrons to the heme Fe(III), one of which moves into the heme ring, forming Fe(IV). The heme ring then removes an electron from Tyr 385. This then removes a H. from the substrate, arachidonic acid. These steps are illustrated in Figure \(37$. The heme site is not shown in the next steps. Next, the reactive dioxygen, itself a ground state diradical, form a bound to the carbon-free radical in arachidonic acid. The rest of the reaction ensues as shown in Figure $38$. Figure $39$ shows an interactive iCn3D model of the mouse cyclooxygenase 2 (5COX) Figure $39$: mouse cyclooxygenase 2 (5COX) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...wt9ECkgsyu4Vs8 Here is a model of human cyclooxygenase 1 structure (6Y3C): https://structure.ncbi.nlm.nih.gov/i...joK7oV63vLSi1A Figure $39$ shows an interactive iCn3D model of the mouse COX-2 with bound arachidonic acid (3KRK) Figure $39$: Mouse COX-2 with bound arachidonic acid (3KRK) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...yAtJnTkTmuNreA The following key binding and catalytic residues are shown and labeled: • Arg-120 and Tyr-355 are close to the carboxylate of AA • Phe205, Phe209, Val228, Val344, Phe381, and Leu534 form a hydrophobic groove for the ω-end of AA. • Ser 530, which is above this, gets acetylated by aspirin • Tyr 385, near C13 in arachidonic acid, forms the free radical which removes a single electron from C13 Prostaglandins, which were first isolated from prostate glands, serve as powerful, but labile local hormones which are mediators of pain, inflammation, and immune and clotting activity. The cyclooxygenase activity is inhibited by aspirin, which probably accounts for most of its anti-inflammatory and analgesic properties. Aspirin, acetylsalicylic acid, acetylates a reactive Ser 530 in the active site. Another nonsteroidal anti-inflammatory drug (NSAID) with similar properties is Ibuprofen (Advil). Acetaminophen (Tylenol) is also considered a member of this drug class, even though it doesn't have anti-inflammatory properties. The question has arisen as to why. It now turns out that there are three different types of COX, I, II, and III. COX III is expressed in the brain and might be involved in pain pathways. Acetaminophen appears to work on this COX, as shown in Table $1$ below (Bazan et al.). Cyclooxygenase Activities COX Expression Function Inhibitors COX 1 constitutively organ pain, platelet function, stomach protection NSAIDs including aspirin COX 2 induced by growth factors, neurotransmitters, inflammatory cytokines, oxidative stress, and injury. Constitutively in the brain, kidney Inducible COX2: inflammation, pain, fever Constitutive COX2: synaptic plasticity NSAIDs, COX 2 inhibitors including celecoxib (Celebrex ) which has few GI problems associated with its use COX 3 constitutively, high in brain, heart pain pathways, not inflammation pathways acetaminophen (no GI problems, great fever reducer), some NSAIDs Fish n-3 fatty acids and health We mentioned the importance of arachidonic acid in signal transduction in the lipid chapter. In addition, the importance of n-3 fatty acids to health was discussed as well. As mentioned above, arachidonic acid is cleaved from the C2 or sn-2 position of membrane phospholipids and modified by cyclooxygenase or lipoxygenase to form prostaglandins and leukotrienes, both potent local biological mediators. Linoleic acid and 22:6n-3 (DHA or docosahexaenoic acid) are also found in membrane phospholipids at the sn-2 position. What is the mechanism for the health-protective effects of n-3 fatty acids like DHA? In human tissue, DHA, 22:6n-3 or 22:6Δ5,8,11,14,17,20 is the most abundant n-3 polyunsaturated fatty acids (PUFAs). Since it is synthesized from linolenic acid (as is EPA), a deficiency of linolenic acid in the diet will lead to lowered levels of 22:6n-3 in tissues, with ensuing health effects. Since these lipids are involved in membrane structure, signal transduction, and hormone synthesis, diverse effects of dietary n-3 PUFA deficiency will be observed. 50% of all fatty acids in the sn1 and sn2 position of membrane phospholipids of rod outer segments (in the retina) are 22:6(n-3). Cognitive dysfunctions (loss of memory, etc.) have been linked to decreased levels of 22:6(n-3) in the brain. This fatty acid binds to retinoid X receptors which then activate (through linked binding reactions) nuclear receptors, leading to alterations in gene transcription in the CNS. In other tissues, 22:6(n-3) rarely exceeds 10% of membrane fatty acids, but this percentage can be increased in cells with increases in a precursor, 20:5(n-3). DHA might affect lipid rafts in the membrane, which would affect the movement of important membrane protein receptors (and associated proteins) in the membrane, altering cell response to environmental stimuli. DHA and EPA affect arachidonic acid conversion to prostaglandins and leukotrienes. EPA binds less tightly to cyclooxygenase I and is a poor substrate for the enzyme, both effects which inhibit the formation of prostaglandins and signaling processes mediated by them. This explains why n-3 fatty acids have anti-inflammatory effects. In addition, n-3 fatty acids have noticeable effects on gene transcription, which remain as long as these fatty acids are present in high levels in the diet These and other fatty acids bind to fatty acid-activated transcription factors called PPARs (peroxisome proliferator receptors - alpha, beta and gamma 1 and 2). These receptors regulate, through alterations in gene expression, proteins involved in lipid metabolism. Other fatty acid-dependent transcription factors are known as well. PPARs bind 20:5(n-3) with a micromolar Kd and change the conformation of the protein to a form than can bind other proteins, ultimately altering gene expression. Table $2$: Some biological effects of n-3 polyunsaturated fatty acids EPA (20:5) and DHA (22:6). Adapted from Jump D. The Biochemistry of n-3 Polyunsaturated fatty acids. J. Biol. Chem. 277, pg 8755 (200) Organ(s) Effect Mechanism acts through central nervous system improve cognitive function membrane composition; retinoic X receptor alpha retina improve acuity membrane composition immune immunosuppressive; anti-inflammatory membrane composition; rafts cardiovascular anti-arrhythmia; anti-clotting membrane composition; rafts; eicosanoids serum lipids lowers triglycerides (a risk factor for cardiovascular. disease) peroxisome proliferator receptor alpha and gamma liver decrease lipid synthesis; increase fatty acid oxid. decrease VLDL synthesis sterol reg. element bind. protein; PPAR alpha PPAR alpha Recognition of Unsaturated Fatty Acids by Membrane G-protein-coupled receptors The positive health effects of the n-3 (ω-3) fatty acids appear to involve those fatty acids acting as hormones that bind to a special membrane receptor called G protein-coupled receptor 120 (GPCR120).  We will discuss GPCRs in greater detail in Chapter 28.2.  When a hormone bounds to the membrane GPCR (a transmembrane protein with 7 membrane-spanning helices), the intracellular domain of the GPC alters how it interacts with a variety of binding partners inside the cell.  The GPCRs are given that name since most interact with heterotrimeric G proteins inside the cell.  These G proteins have a β, γ, and a variety of different α subunits (αq, αi, αs).  Activation of different α subunits leads to different downstream effects as part of an elaborate signaling system in the cell.  When the health-promoting polyunsaturated n-3 (ω-3) fatty bind to GPCR120, here are some of the beneficial health effects on metabolism, as outlined in Table $3$ below. Protein involved in GPCR120 signaling Downstream effect q Increases Ca2+ levels in the cell, which act as a second signal (messenger) Increases GLUT4 (glucose transporter) translocation to the cell membrane; Increases secretion of glucagon-like peptide 1 (GLP1) i Increases insulin secretion Inhibits Ghrelin (hunger hormone) secretion s  Controls fat synthesis non G protein GPCR kinase (GRK)/β-arrestin 2 Inhibits the NLRP3 inflammasome and is ant-iinflammatory The data in the table above is summarized in Figure $40$ below. Figure $40$: Schematic overview of G protein–and arrestin–mediated GPR120 signaling and related functions.  Chunyou Mao et al. Science 380, eadd6220 (2023). https://www.science.org/doi/10.1126/science.add6220.  With permission from the AAAS. GPCR120 and differentiate fatty acids based on the number and position of the double bond in the acyl chain of the ligand.  Even though saturated fatty acids can bind to GPCR 120, only the n-3 (ω-3) fatty acids elicit the health effects. CryoEM structures show the fatty acids buried in an L-shaped conformation in the GPCR.  The protein engages in noncovalent interactions, specifically π:π interactions, with the double bonds in acyl chain. These interactions, which depend on the number of double bonds in the n-3 (ω-3) fatty acid differentially biases the GPCR towards the promotion of the specific effects for each different fatty acid. Figure $41$ shows an interactive iCn3D model of the eicosapentaenoic acid bound GPR120-Gi complex (8ID9). Figure $41$: Eicosapentaenoic acid bound GPR120-Gi complex (8ID9). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...vgMGJRTr3emNj7 The color coding is as follows: • GPCR120 - cyan (the membrane would be perpendicular across the 7 membrane helices) with the bound EPA • Gβ - magenta • Gγ - blue • EPA -spacefill Figure $42$ shows an interactive iCn3D model showing the nonpolar environment of eicosapentaenoic acid bound to  GPR120 (8ID9). Figure $42$: Nonpolar environment of eicosapentaenoic acid bound to  GPR120 (8ID9). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...SszD1K3B9ieND7 Dark green shows the hydrophobic amino acids

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/21%3A_Lipid_Biosynthesis/21.01%3A_Biosynthesis_of_Fatty_Acids_and_Eicosanoids.txt

Search Fundamentals of Biochemistry by William (Bill) W. Christie and Henry Jakubowski. This section is an abbreviated and modified version of material from the Lipid Web, an introduction to the chemistry and biochemistry of individual lipid classes, written by William Christie. Introduction All eukaryotic organisms and even a few prokaryotes can synthesize triacylglycerols, and in animals, many cell types and organs have this ability, but the liver, intestines, and adipose tissue are most active with most of the body stores in the last of these. Within all cell types, even those of the brain, triacylglycerols are stored as cytoplasmic 'lipid droplets' enclosed by a monolayer of phospholipids and hydrophobic proteins such as the perilipins in adipose tissue or oleosins in seeds. These lipid droplets are now treated as distinctive organelles, with their own characteristic metabolic pathways and associated enzymes - no longer boring blobs of fat. However, they are not unique to animals and plants, as Mycobacteria and yeasts have similar lipid inclusions. The lipid serves as a store of fatty acids for energy, which can be released rapidly on demand, and as a reserve of fatty acids for structural purposes or as precursors for eicosanoids. In addition, lipid droplets serve as a protective agency in cells to sequester any excess of biologically active and potentially harmful lipids such as free fatty acids, oxylipins, diacylglycerols, cholesterol (as cholesterol esters), retinol esters, and coenzyme A esters. While triacylglycerols are essential for normal physiology, an excessive accumulation in human adipose tissue and other organs results in obesity and other health problems, including insulin resistance, steatohepatitis, and cardiomyopathy. Accordingly, there is considerable pharmaceutical interest in drugs that affect triacylglycerol biosynthesis and metabolism. Biosynthesis of Triacylglycerols Three main pathways for triacylglycerol biosynthesis include the sn-glycerol-3-phosphate and dihydroxyacetone phosphate pathways, which predominate in liver and adipose tissue, and a monoacylglycerol pathway in the intestines. In maturing plant seeds and some animal tissues, a fourth pathway has been recognized in which a diacylglycerol transferase is involved. The most important route to triacylglycerols is the sn-glycerol-3-phosphate or Kennedy pathway, first described by Eugene Kennedy and colleagues in the 1950s, from which more than 90% of liver triacylglycerols are produced. Figure \(1\) shows the pathway from glycerol-3-phosphate to triacylglycerols (the Kennedy pathway) and some preceding reactions that generate glycerol-3-phosphate. In this pathway, the main source of the glycerol backbone has long been believed to be sn-glycerol-3-phosphate produced by the catabolism of glucose (glycolysis) or to a lesser extent by the action of the enzyme glycerol kinase on free glycerol. However, there is increasing evidence that a significant proportion of glycerol is produced de novo by a process known as glyceroneogenesis via pyruvate. Indeed, this may be the main source in adipose tissue. Subsequent reactions occur primarily in or at the endoplasmic reticulum. First, the precursor sn-glycerol-3-phosphate is esterified by a fatty acid coenzyme A ester in a reaction catalyzed by a glycerol-3-phosphate acyltransferase (GPAT) at position sn-1 to form lysophosphatidic acid, and this is in turn acylated by an acylglycerophosphate acyltransferase (AGPAT) in position sn-2 to form a key intermediate in the biosynthesis of all glycerolipids - phosphatidic acid. Numerous isoforms of these enzymes are known; they are expressed with specific tissue and membrane distributions, and they are regulated in different ways. Let's look at some of the enzymes involved in some key reactions in the synthesis of triacylglycerols. GPAT Enzymes Glycerol-3-phosphate acyltransferase (GPAT) catalyzes the first step in the pathway in most tissue. From an enzymatic perspective, it can be considered rate-limiting for the pathways since its specific activity is slow. In mammals, there are four isoforms (GPAT1 in the outer membrane of the mitochondria, GPAT2 also in the mitochondria, and GPAT3 and 4, both in the ER. The activity of GPAT1 is activated by insulin, presumably through phosphorylation) and inhibited by AMP-activated protein kinase (AMPK). Fatty acyl-CoA in the outer membrane could be used for TAG synthesis through the Kennedy pathway in times of energy abundance or imported into the matrix through the carnitine-acyl-CoA cycle in times of energy need. Overexpression of GPAT1 in liver cells increases TAG synthesis and decreases β-oxidation. GPAT1 deficient mice in contrast have higher levels of β-oxidation. Figure \(2\) shows an interactive iCn3D model of the AlphaFold predicted structure of human mitochondrial glycerol-3-phosphate acyltransferase 1 - GPAT1 (Q9HCL2). Figure \(2\): AlphaFold predicted structure ofhuman mitochondrial glycerol-3-phosphate acyltransferase 1 - GPAT1 (Q9HCL2). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...kuvBTPo5mNK8p9 Four conserved motifs involved in catalysis (red and cyan) and binding (orange and yellow) are shown. Significant sections are still disordered in the model. Two highly conserved amino acids, His 230 and Asp 235 in the red catalytic motif are shown in CPK-colored sticks and labeled. The black helices are predicted outer mitochondrial membrane transmembrane helices. Both the N- and C-terminal amino acids project into the cytoplasm. Phosphatidic acid phosphohydrolases (PAPs or ‘phosphatidate phosphatases’ or ‘lipid phosphate phosphatases’). PAPs are also important as they produce sn-1,2-diacylglycerols as essential intermediates in the biosynthesis not only of triacylglycerols but also of phosphatidylcholine and phosphatidylethanolamine (and of monogalactosyldiacylglycerols in plants). This is a key branch-point in lipid biosynthesis as it may dictate the flow of lipids for storage or membrane biogenesis. Much of this phosphatase activity leading to triacylglycerol biosynthesis in animals reside in three related cytoplasmic proteins, termed lipins, i.e., lipin-1, lipin-2 and lipin-3, which have tissue-specific roles in glycerolipid synthesis. Unusually, these were characterized and named before the nature of their enzymatic activities was determined. Each of the lipins appears to have distinctive expression and functions, but lipin-1 (PAP1) in three isoforms (designated 1α, 1β, and 1γ) accounts for most of the PAP activity in adipose tissue and skeletal muscle in humans. Lipin 2 is the most abundant lipin in the liver but is also expressed substantially in the small intestine, macrophages, and some regions of the brain, while lipin 3 activity overlaps with that of lipin 1 and lipin 2 and is found in the gastrointestinal tract and liver. Lipins are cytosolic enzymes but associate transiently with membranes to access their substrate, i.e., they are translocated to the endoplasmic reticulum in response to elevated levels of fatty acids within cells, although they do not have trans-membrane domains. Lipin-1 activity requires Mg2+ ions and is inhibited by N-ethylmaleimide, whereas the membrane-bound activity responsible for synthesizing diacylglycerols as a phospholipid intermediate is independent of Mg2+ concentration and is not sensitive to the inhibitor. Perhaps surprisingly, lipin-1 has a dual role in that it operates in collaboration with known nuclear receptors as a transcriptional coactivator to modulate lipid metabolism (lipin 1α) while lipin 1β is associated with the induction of lipogenic genes such as fatty acid synthase, stearoyl-CoA desaturase, and DGAT. They can have profound effects on signaling in a variety of cell types. Abnormalities in lipin-1 expression are known to be involved in some human disease states that may lead to metabolic syndrome and inflammatory disorders. Lipin 2 is a similar phosphatidate phosphohydrolase, which is regulated dynamically by fasting and obesity (in mice). PAP depend on divalent Mg2+ ions and are sensitive to N-ethylmaleimide. Lipin has a N-Lip (N-terminal) and C-Lip (C-terminal) domain. The crystal structure of lipin with a 250 amino acid regions between the two domains removed has been solved. The domain structures of humans, the yeast Saccharomyces cerevisiae (SC) and its truncated form (Tt-Pah2), and the structure of the Tt-Pah2 truncated fragment of the enzyme from Tetrahymena thermophila, are shown in Figure \(3\). Panel (a) shows the PAP-catalyzed dephosphorylation reaction. Panel (b) shows the domain architecture of PAPs drawn to scale. The positions of the nuclear localization signal (NLS), conserved Trp-motif (purple W), catalytic DxDxT motif, and fatty liver dystrophy (fld2J) mutation are indicated. Panel (c) shows the the wild-type (WT) Tt Pah2 is catalytically active and the D146A mutant of the DxDxT motif eliminates activity. Panel (d) shows the overall structure of Tt Pah2, which contains an immunoglobulin-like (Ig-like) domain and a HAD-like catalytic domain. The Ig-like domain is formed by the N-Lip (cyan) and C-Lip (pink) regions connected by a short linker (gray loop) that would be replaced by the extended 500-residue linker in human lipins and a 250-residue linker in Sc Pah1. A calcium ion (Ca2+, yellow sphere) is bound in the active site of the HAD-like catalytic domain. The top view (right) of the enzyme shows the N-Lip co-folding with the C-Lip to form the Ig-like domain. Figure \(4\)s shows a closeup of the active site of Tt Pah2 and a mechanism for the hydrolytic removal of the phosphate on DAG. Figure \(5\) shows an interactive iCn3D model of the Tetrahymena Thermophila lipin phosphatidic acid phosphatase with magnesium (6TZZ). Figure \(5\): Tetrahymena Thermophila lipin phosphatidic acid phosphatase with magnesium (6TZZ). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...GTusCCDP5eRrt7 The N-terminal domain is shown in lavender and the C-terminal domain is in gray. Key amino acids in the active site are shown in CPK-colored sticks and labeled. Diacylglycerol acyltransferases (DGAT) In the final step in this pathway, the 1,2-diacyl-sn-glycerol intermediate is acylated by diacylglycerol acyltransferases (DGAT), which can utilize a wide range of fatty acyl-CoA esters to form the triacyl-sn-glycerol. There are two DGAT enzymes, which are structurally and functionally distinct. In animals, DGAT1 is located mainly in the endoplasmic reticulum and is expressed in skeletal muscle, skin, and intestine, with lower levels of expression in the liver and adipose tissue. It is believed to have dual topology contributing to triacylglycerol synthesis on both sides of the membrane of the endoplasmic reticulum but esterifying only pre-formed fatty acids of exogenous origin. Perhaps surprisingly, DGAT1 is the only one present in the epithelial cells that synthesize milk fat in the mammary gland. Also, DGAT1 can utilize a wider range of substrates, including monoacylglycerols, long-chain alcohols (for wax synthesis), and retinol, and it is reported to have an important role in protecting the endoplasmic reticulum from the lipotoxic effects of high-fat diets. Orthologs of this enzyme are present in most eukaryotes, other than yeasts, and they are especially important in plants. Figure \(6\) shows an interactive iCn3D model of Human Diacylglycerol Acyltransferase 1 in complex with oleoyl-CoA (6VP0) Figure \(6\): Human Diacylglycerol Acyltransferase 1 in complex with oleoyl-CoA (6VP0) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...adyWg9fy2hh8r8 The red layer is the extracellular leaflet and the blue is the intracellular leaflet. The protein is a dimer with one subunit shown in gray and one in light. A substrate, oleoyl-CoA, is shown in spacefill with CPK colors in both monomers. The acyl chain with a clear kink induced by the double bond in the oleoyl acyl group is clearly in the hydrophobic interior of the bilayer. The gray monomer shows the atoms within 4 Å from the oleoly-CoA is shown as surface color-coded by hydrophobicity with the green most hydrophobic. The active site His 415 and Met 434 which hydrogen bonds to it are shown in the light cyan subunit in sticks, CPK colors, and labeled. Three other conserved polar side chains, Asn378, Gln437, and Gln465, which are in the active site region, are also shown. Each monomer in the homodimer has 9 transmembrane helices with 8 forming a unique fold called the MBOAT which encloses a chamber with catalytic side chains. The substrates, fatty acyl-CoA, and DAG have different entrances to the chamber with the acyl-CoA occupying a cytosolic tunnel. The cytosolic face interacts with the CoA and the acyl chain extends into the ER membrane. The channel appears bent which probably allows the acylation of DAGs and long-chain alcohols, but not cholesterol, a rigid, planar molecule that is acylated to form an ester by a different enzyme, acyl CoA:cholesterol acyltransferases. Hence it appears that DAG, the acyl-acceptor, binds in a hydrophobic tunnel from within the bilayer and the acyl-CoA binds in another tunnel, allowing the transfer of the acyl group. DGAT2 is the main form of the enzyme in hepatocytes and adipocytes (lipid droplets), although it is expressed much more widely in tissues. It is associated with distinct regions of the endoplasmic reticulum, at the surface of lipid droplets, and in mitochondria, and it esterifies fatty acids of both endogenous and exogenous origin. DGAT2 is believed to have a targeting domain that enables it to tether between the endoplasmic reticulum and lipid droplet thereby channeling triacylglycerols from the synthesis site in the endoplasmic reticulum to the nascent lipid droplet, where they accumulate and lead to the expansion of the latter (see below). Both enzymes are important modulators of energy metabolism, although DGAT2 appears to be especially important in controlling the homeostasis of triacylglycerols in vivo. As the glycerol-3-phosphate acyltransferase (GPAT) has the lowest specific activity of these enzymes, this step may be the rate-limiting one. However, DGATs are the dedicated triacylglycerol-forming enzymes, and they are seen as the best target for pharmaceutical intervention in obesity and attendant ailments; clinical studies of DGAT1 inhibitors are at an early stage. In a second pathway for triacylglycerol biosynthesis, dihydroxyacetone-phosphate in peroxisomes or endoplasmic reticulum can be acylated with fatty acid CoA esters by a specific acyltransferase to form 1-acyl dihydroxyacetone-phosphate, which is reduced by dihydroxyacetone-phosphate oxidoreductase to lysophosphatidic acid; this can then enter the pathway above to triacylglycerols. The precursor dihydroxyacetone-phosphate is important also as part of the biosynthetic route to plasmalogens, and neutral plasmalogens can be significant components of cytoplasmic droplets in many mammalian cells types but not in adipose tissue. Figure \(7\): Biosynthesis of triacylglycerols from dihydroxyacetone-phosphate In prokaryotes, the glycerol-3-phosphate pathway of triacylglycerol biosynthesis only occurs, but in yeast, both glycerol-3-phosphate and dihydroxyacetone-phosphate can be the primary precursors and synthesis takes place in cytoplasmic lipid droplets and the endoplasmic reticulum. In plants, the glycerol-3-phosphate pathway is most important. In the enterocytes of the intestines after a meal, up to 75% of the triacylglycerols are formed via a monoacylglycerol pathway. In this, 2-monoacyl-sn-glycerols and free fatty acids released from dietary triacylglycerols by the action of pancreatic lipase within the intestines (see below) are taken up by the enterocytes. There, the monoacylglycerols are first acylated by an acyl-coenzyme A:monoacylglycerol acyltransferase with the formation of sn-1,2-diacylglycerols mainly as the first intermediate in the process, though some sn-2,3-diacylglycerols (~10%) are produced. In addition, 1-monoacylglycerols can be synthesized by acylation of glycerol for further acylation. There are three isoforms of the monoacylglycerol acyltransferase in humans of which MGAT2 is most active in the intestines, but also in the liver where an appreciable proportion of the triacylglycerols are formed by the monoacylglycerol pathway, while MGAT1 functions in adipose tissue; the role of MGAT3 is not clear. Finally, the acyl-coenzyme A:diacylglycerol acyltransferase (DGAT1) reacts with the sn-1,2-diacylglycerols (not the sn-2,3 form) to produce triacylglycerols (DGAT1 can also acylate monoacylglycerols). Figure \(8\) shows the biosynthesis of triacylglycerols from monoacylglycerols. Other pathways: In a fourth biosynthetic pathway, which is less well known, triacylglycerols are synthesized by a transacylation reaction between two racemic diacylglycerols that are independent of acyl-CoA. The reaction was first detected in the endoplasmic reticulum of intestinal micro villus cells and is catalyzed by a diacylglycerol transacylase. Both diacylglycerol enantiomers participate in the reaction with equal facility to transfer a fatty acyl group with the formation of triacylglycerols and a 2-monoacyl-sn-glycerol. A similar reaction has been observed in seed oils. Figure \(9\): Biosynthesis of triacylglycerols from diacylglycerol transacylases It has been suggested that this enzyme may function in remodeling triacylglycerols post-synthesis, especially in oil seeds, and it may be involved in similar processes in the liver and adipose tissue, where extensive hydrolysis/re-esterification is known to occur. There is evidence for selectivity in the biosynthesis of different molecular species in a variety of tissues and organisms, which may be a consequence of the varying biosynthetic pathways. Also in adipose tissue, fatty acids synthesized de novo are utilized in different ways from those from external sources in that they enter positions sn-1 and 2 predominantly, while a high proportion of the oleic acid synthesized in the tissue by desaturation of exogenous stearic acid is esterified to position sn-3. Among other potential routes to the various intermediates, lysophosphatidic acid and phosphatidic acid can be synthesized in mitochondria, but they must then be transported to the endoplasmic reticulum before they enter the pathway for triacylglycerol production. 1,2‑Diacyl-sn-glycerols are also produced by the action of phospholipase C on phospholipids and can be utilized for triacylglycerol biosynthesis. In the glycerol-3-phosphate and other pathways, the starting material is of defined stereochemistry and each of the enzymes catalyzing the various steps in the process is distinctive and can have preferences for particular fatty acids (as their coenzyme A esters) and for particular fatty acid combinations in the partially acylated intermediates. It should not be surprising, therefore, that natural triacylglycerols exist in enantiomeric forms with each position of the sn-glycerol moiety esterified by different fatty acids. Triacylglycerol Metabolism in the Intestines, Liver, and Mammary Gland Fat comprises up to 40% of the energy intake in the human diet in Western countries, and a high proportion of this is in the form of triacylglycerols. The process of fat digestion is begun in the stomach with acid-stable gastric or lingual lipases, the extent of which depends on species but may be important for efficient emulsification. However, this is insignificant in quantitative terms in comparison to the reaction with pancreatic lipase, which occurs in the duodenum. Entry of triacylglycerol degradation products into the duodenum stimulates the synthesis of the hormone cholecystokinin and causes the gall bladder to release bile acids, which are strong detergents and act to emulsify the hydrophobic triacylglycerols so increasing the available surface area. In turn, cholecystokinin stimulates the release of the hydrolytic enzyme pancreatic lipase together with a co-lipase, which is essential for the activity of the enzyme. Pancreatic lipase, co-lipase, bile salts, and calcium ions act together in a complex at the surface of the emulsified fat droplets to hydrolyze the triacylglycerols. The process is regiospecific and results in the release of the fatty acids from the 1 and 3 positions and the formation of 2-monoacyl-sn-glycerols as shown in Figure \(10\). Spontaneous isomerization of the latter to 1(3)-monoacyl-sn-glycerols occurs to some extent, and these can be degraded completely by the enzyme to glycerol and free fatty acids. Other lipases hydrolyze the phospholipids and other complex lipids in foods at the same time. This process is somewhat different in neonates and young infants, in whom pancreatic lipase is less active but is effectively replaced by lipases in breast milk and by an acid gastric lipase (pH optimum 4-6). There is evidence that the regiospecific structure of dietary triacylglycerols affects the uptake of particular fatty acids and may influence further lipid metabolism in humans. Incorporation of palmitic acid into the position sn-2 of milk fat may be of benefit to the human infant (as a source of energy for growth and development), although it increases the atherogenic potential for adults. In addition, 2-monoacylglycerols and 2-oleoylglycerol especially have a signaling function in the intestines by activating a specific G‑protein coupled receptor GPR119, sometimes termed the ‘fat sensor’. When stimulated, this causes a reduction in food intake and body weight gain in rats and regulates glucose-stimulated insulin secretion. The free fatty acids released have a similar effect, though by a very different mechanism, via the receptor GPR40. Overall, it has become evident that triacylglycerol metabolism in the intestine has regulatory effects on the secretion of gut hormones and subsequently on systemic lipid metabolism and energy balance. The free fatty acids and 2-monoacyl-sn-glycerols are rapidly taken up by the intestinal cells, from the distal duodenum to the jejunum, via specific carrier molecules but also by passive diffusion. A specific fatty acid binding protein prevents a potentially toxic build-up of unesterified fatty acids and targets them for triacylglycerol biosynthesis. The long-chain fatty acids are converted to the CoA esters and esterified into triacylglycerols by the monoacylglycerol pathway as described above. In contrast, short and medium-chain fatty acids (C12 and below) are absorbed in unesterified form and pass directly into the portal blood stream, where they are transported to the liver to be oxidized. Subsequently, the triacylglycerols are incorporated into lipoprotein complexes termed chylomicrons in the enterocytes in the small intestines. In brief, these consist of a core of triacylglycerols together with some cholesterol esters that is stabilized and rendered compatible with an aqueous environment by a surface film consisting of phospholipids, free cholesterol, and one molecule of a truncated form of apoprotein B (called apo B48). A cartoon structure of a chylomicron is shown in Figure \(11\). These particles are secreted into the lymph and thence into the plasma for transport to the peripheral tissues for storage or structural purposes. Adipose tissue in particular exports appreciable amounts of the enzyme lipoprotein lipase, which binds to the luminal membrane of endothelial cells facing into the blood, where it rapidly hydrolyses the passing triacylglycerols at the cell surface releasing free fatty acids, most of which are absorbed into the adjacent adipocytes and re-utilized for triacylglycerol synthesis within the cell. The chylomicrons remnants eventually reach the liver, where the remaining lipids are hydrolyzed at the external membranes by a hepatic lipase and absorbed. The fatty acids within the liver can be utilized for a variety of purposes, from oxidation to the synthesis of structural lipids, but a proportion is re-converted into triacylglycerols, and some of this is stored as lipid droplets within the cytoplasm of the cells (see next section). In addition, phosphatidylcholine from the high-density lipoproteins is taken up by the liver, and a high proportion of this is eventually converted to triacylglycerols. In a healthy liver, the levels of triacylglycerols are low (<5% of the total lipids), because the rates of acquisition of fatty acid from plasma and synthesis de novo within the liver are balanced by rates of oxidation and secretion into plasma. On the other hand, excessive accumulation of storage triacylglycerols is associated with fatty liver, insulin resistance, and type 2 diabetes. Most of the newly synthesized triacylglycerols are exported into the plasma in the form of very-low-density lipoproteins (VLDL), consisting again of a triacylglycerol and cholesterol ester core, surrounded by phospholipids and free cholesterol, together with one molecule of full-length apoprotein B (100 kDa), apoprotein C and sometimes apoprotein E. These particles in turn are transported to the peripheral tissues, where they are hydrolyzed and the free acids absorbed. Eventually, the remnants are returned to the liver. In the mammary gland, triacylglycerols are synthesized in the endoplasmic reticulum and large lipid droplets are produced with a monolayer of phospholipids derived from this membrane. These are transported to the plasma membrane and bud off into the milk with an envelope comprised of the phospholipid membrane to form milk fat globules as food for the newborn. The process is thus very different from that involved in the secretion of triacylglycerol-rich lipoproteins from other organs. Triacylglycerol Synthesis and Catabolism (Lipolysis) in Adipocytes and Lipid Droplets Adipose tissue and the adipocytes are characterized by accumulations of triacylglycerols, which act as the main energy store for animals, although they also cushion and insulate the body. Large fat depots occur around internal organs such as the liver, and also subcutaneously, and each of these may react differently to metabolic constraints. Thus, triacylglycerols stored when there is a surplus of nutrients are mobilized for energy production during starvation. Adipose tissue also functions as a reserve of bioactive lipids, such as eicosanoids and lipid-soluble vitamins, and when required provides structural components, including fatty acids, cholesterol, and retinol, for membrane synthesis and repair. By buffering against fatty acid accumulation that might exceed their capacity, non-adipose cells defend themselves in this way against lipotoxicity while providing a rapid source of energy and essential metabolites by sensing and responding rapidly to changes in systemic energy balance. Brown and beige fat have special properties and are discussed below, while bone marrow adipocytes (70% of the available space) have distinctive functions also. Similarly, within most other animal cells, including most cell types in the brain, a proportion of the fatty acids taken up from the circulation is converted to triacylglycerols as described above and incorporated into cytoplasmic lipid droplets (also termed 'fat globules', 'oil bodies', 'lipid particles' or 'adiposomes'). In adipocytes, the lipid droplets can range from up to 200 μm in diameter, while other cell types contain smaller lipid droplets of the order of 50 nm in diameter. The triacylglycerol droplets together with cholesterol esters and other neutral lipids are surrounded by a protective monolayer that includes phospholipids, cholesterol, and hydrophobic proteins. The phospholipid component of the monolayer consists mainly of phosphatidylcholine and phosphatidylethanolamine derived from cytosolic leaflets of the endoplasmic reticulum and plasma membrane. Among the proteins are many that function directly in lipid metabolism, and they include acyltransferases, lipases, perilipins, caveolins, and the Adipose Differentiation Related Protein (ADRP or adipophilin). Acting in concert with other cellular organelles, they function in many different metabolic processes facilitating coordination and communication between different organelles and acting as vital hubs of cellular metabolism. Cytosolic lipid droplets with similar metabolic activities are found in most eukaryotic cells, including those of the fruit fly Drosophila melanogaster, and many aspects of triacylglycerol processing and regulation parallel those in humans. They are also present in the cytoplasm of some prokaryotes and in the plastids and other organelles of plants (see below). Lipid droplet assembly This process takes place in sub-domains of the endoplasmic reticulum, where at least one isoform of each of the enzymes of triacylglycerol biosynthesis, from acyl-CoA synthetases through to glycerol-3-phosphate acyltransferases, is located probably in a protein assembly or 'interactome'. As triacylglycerols accumulate, they reach a critical level when a spontaneous condensation or nucleation by phase separation occurs, leading to the formation of an oil blister within the hydrophobic bilayer region so attracting perilipins and other proteins that allow lipid droplets to grow further in patches of the membrane as lens-like swellings between the two membrane leaflets. The protein seipin stabilizes the nascent droplets with minimal disruption to the membrane and enables them to mature. Seipin monomers assemble into a decameric cage-like structure and sit in the ER to provide a space that permits triacylglycerol molecules to interact with each other, rather than with phospholipid acyl chains, a process that is probably aided by trans-membrane protein segments. This enables phase separation of the triacylglycerols, lens formation, and growth to a point where the seipin oligomer opens toward the cytoplasm so the lens can form a budding lipid droplet. Figure \(12\) presents a cartoon that shows the role of ER membrane seipins and other proteins in the assembly of lipid droplets. In panel (A), seipin is enriched at the endoplasmic reticulum (ER)/lipid droplet (LD) contact sites and is crucial in triglycerides (TAG) flow from the ER to the LD. Seipin interacts with Perilipin1 (PLIN1) and with several TAG synthetic enzymes such as glycerol-3-phosphate acyltransferase (GPAT3), 1-acyl-sn-glycerol-3-phosphate acyltransferase beta (AGPAT2), and LIPIN. No interaction with diacylglycerol acyltransferases (DGAT) has been formally reported. Magré J et al, Int J Mol Sci. 2022 Jan 11;23(2):740. doi: 10.3390/ijms23020740. Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/ In panel (B), representing the fasting state, seipin is enriched at the ER/mitochondria (Mt) contact sites, also named mitochondria-associated membranes (MAM) and participates in ER/mitochondria calcium (Ca2+) flux and mitochondrial activity. Seipin is near MAM Ca2+ regulators IP3(inositol 1,4,5-trisphosphate) receptor (IP3R), voltage-dependent anion channel (VDAC), and sarco-/ER Ca2+ ATPase 2 (SERCA2) as well as glycerol-3-phosphate (G3P), lysophosphatidic acid (LPA), phosphatidic acid (PA), and diacylglycerol (DAG). Figure \(13\) shows an interactive iCn3D model of the S. cerevisiae seipin Homo 10-mer flexible cage at lipid droplet formation sites (7RSL). Figure \(13\): S. cerevisiae seipin homo 10-mer flexible cage at lipid droplet formation sites (7RSL). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...BjYLiQXVJjBpk8 The N-terminal transmembrane segment (TMS) on each monomer is shown in cyan while the C-terminal TMS is shown in magenta. These form the cage sides and top and anchor the complex in the ER membrane. The side chains of a conserved switch sequence (F232xxGLR) on each monomer are shown in spacefill, CPK colors. The gray ribbons represent the part of the complex that is in the lumen of the ER. Adjacent monomers in the homo 10-mer cage, even though they are identical in sequences, adopt different orientations for two helices that enable a binary switch in the conformation of the complex. Two adjacent subunits, the A chain in blue and the B chain in orange, are shown, along with their overlap, in Figure \(14\). A highly conserved "switch" region is shown in green with its highly conserved motif F232xxGLR shown as sticks with CPK colors. Note the large conformational change in the switch region which occurs with a large conformational change of the two helices. This suggests that there are two overall conformations of the complex, a closed form allowing accumulation of TAGs and an open form which allows the budding and release of lipid droplets. A possible model for the formation and budding of lipid droplets is illustrated in Figure \(15\). As in the above figure, the A (open) chain is shown in blue while the B (closed) chain is shown in orange. Some of the surface proteins on lipid droplets can extend long helical hairpins of hydrophobic peptides deep into the lipid core. For example, perilipins constitute a family of at least five phosphorylated proteins that bind to droplets in animals and share a common region, the so-called ‘PAT’ domain, named for the three original members of the family that include perilipin and ADRP. Proteins related evolutionarily to these are found in more primitive organisms, including insects, slime molds, and fungi, but not in the nematode Caenorhabditis elegans. In mammals, perilipin A (or 'PLIN1' or more accurately the splice variant 'PLIN1a') is a well-established regulator of lipolysis in adipocytes, and it is believed to be involved in the formation of the large lipid droplets in white adipose tissue. The perilipins PLIN1 and PLIN2 have functions in triacylglycerol metabolism in tissues other than adipocytes, and PLIN2 is the main perilipin in hepatocytes; PLIN5 operates in tissues that oxidize fatty acids such as the heart. Other surface proteins of lipid droplets are enzymes intimately involved in triacylglycerol metabolism, although there is a suggestion that cytoplasmic droplets may act as a storage organelle for hydrophobic proteins whose function is elsewhere in the cell. Lipolysis When fatty acids are required by other tissues for energy or other purposes, they are released from the triacylglycerols by the sequential actions of three cytosolic enzymes at neutral pH, i.e., adipose triacylglycerol lipase (ATGL), hormone-sensitive lipase (HSL) and monoacylglycerol lipase, which cycle between the cytoplasmic surfaces of the endoplasmic reticulum and the surface layer of lipid droplets. Simplistically, ATGL hydrolyses triacylglycerols to diacylglycerols, which are hydrolyzed by HSL to monoacylglycerols before these are hydrolyzed by the monoacylglycerol lipase to complete the process. Lipolysis proceeds in a highly ordered manner with stimulation through cell-surface receptors via neurotransmitters, hormones, and autocrine/paracrine factors that activate various intracellular signaling pathways and increase kinase activity. A protein perilipin (PLIN1) has been described as "the gatekeeper of the adipocyte lipid storehouse" that regulates lipolysis by acting as a barrier to lipolysis in non-stimulated cells. However, on β-adrenergic stimulation during fasting, it is phosphorylated by the cAMP-protein kinase, which changes its shape and reduces its hydrophobicity, and in the process activates lipolysis. An isoform, perilipin A, is the main regulatory factor in white adipose tissue. However, many other proteins interact with the three enzymes to modulate their activity, location, and stability. The adipose triacylglycerol lipase, which initiates the process, was discovered surprisingly recently. It is structurally related to the plant acyl-hydrolases in that it has a patatin-like domain in the NH2-terminal region (patatin is a non-specific acyl-hydrolase in potatoes). Specific transport mechanisms guide ATGL from the endoplasmic reticulum membrane to lipid droplets, where it is located on the surface both in the basal and activated states. This lipase is specific for triacylglycerols containing long-chain fatty acids, preferentially cleaving ester bonds in the sn-1 or sn-2 position (but not sn-3), and it yields diacylglycerols and free fatty acids as the main products, with low activity only towards diacylglycerols, and none to monoacylglycerols and cholesterol esters. However, it also has transacylase and phospholipase activities, and it hydrolyses retinol esters in hepatic stellate cells. Adipose triacylglycerol lipase can be activated at the same time as hormone-sensitive lipase and is now believed to be rate-limiting for the first step in triacylglycerol hydrolysis. Figure \(16\) shows the sequential hydrolysis of TAGs in adipocytes and lipid droplets. Regulation of the enzymatic activity is a complex process, and for example, a lipid droplet protein designated Gene identification-58 (CGI-58 or ABHD5), is known to be an important activating factor and is required for hydrolysis of fatty acids from position sn-1. In the resting state, this protein binds to perilipin (PLIN1), but on hormonal stimulation, the latter is phosphorylated leading to dissociation and interaction of CGI-58 with phosphorylated ATGL to commence the first step in triacylglycerol hydrolysis. Mutations in adipose triacylglycerol lipase or CGI-58 are believed to be responsible for a syndrome in humans known as ‘neutral lipid storage disease’. A second protein (G0S2) inhibits the enzyme. Hormone-sensitive lipase in various isoforms is a structurally unique member of the large Ser-lipase/esterase family of enzymes in animals. It is regulated by the action of the hormones insulin and noradrenaline by a mechanism that ultimately involves phosphorylation of the enzyme by cAMP-protein kinase (as with perilipin), thereby increasing its activity and causing it to translocate from the cytosol to the lipid droplet to initiate the second step in hydrolysis. Its activity is regulated further by a variety of proteins that include PLINs and fatty acid binding proteins (FABP). Hormone-sensitive lipase has a broad substrate specificity compared to other neutral lipases, and in addition to its activity towards triacylglycerols, it will rapidly hydrolyze diacylglycerols, monoacylglycerols, retinol esters, and cholesterol esters. Diacylglycerols are hydrolyzed ten times as rapidly as triacylglycerols. Within the triacylglycerol molecule, hormone-sensitive lipase preferentially hydrolyses ester bonds in the sn-1 and sn-3 positions, leaving free acids and 2‑monoacylglycerols as the main end products. The monoacylglycerol lipase is believed to be the rate-limiting enzyme in lipolysis, i.e., the final step in triacylglycerol catabolism releasing free glycerol and fatty acids, and it is found in the cytoplasm, the plasma membrane, and in lipid droplets. It is specific for monoacylglycerols, but with no positional specificity, and has no activity against di- or triacylglycerols. As it is the enzyme mainly responsible for the deactivation of the endocannabinoid 2-arachidonylglycerol and is highly active in malignant cancers, it is attracting pharmaceutical interest. A further enzyme, α/β hydrolase containing-6 (ABHD6), is located on the inner leaflet of the plasma membrane and preferentially hydrolyses fatty acids at the sn-1 position over the sn-2 position of monoacylglycerols, and it also hydrolyses lysophospholipids and bis(monoacylglycerol)-phosphate. This has been associated with the development of insulin resistance and the progression of cancer. Additional lipolytic enzymes, including carboxyesterases, are believed to operate against triacylglycerols in cytoplasmic lipid droplets in the liver. Unesterified fatty acids released by the combined action of these three lipases are exported into the plasma for transport to other tissues in the form of albumin complexes, while the glycerol released is transported to the liver for metabolism by either glycolysis or gluconeogenesis. Eventually, the whole organelle can disappear, including the proteins, when they undergo a process of autophagy (lipophagy), i.e., the delivery of the organelles to lytic compartments for degradation. This can occur through direct lysosomal invagination or more often by a multistep process involving the formation of double-membrane vesicles termed 'autophagosomes' around droplets with subsequent lysosomal fusion and degradation of the triacylglycerols by the lysosomal acid lipase. This process is important for the regulation of cellular lipid levels in various tissues and disease conditions, especially during starvation. It is relevant to tumorigenesis and cancer metastasis, and to neurodegenerative and neuroinflammatory diseases. While lipophagy is mechanistically distinct from lipolysis, there is cross-talk between the two. The structure of lipases Lipases carry out hydrolysis reactions (nucleophilic substitution with water as the nucleophile), alcoholysis reactions (with an ROH as the nucleophile), and even transesterification reactions with other fatty acid esters. Lipases are critical enzymes yet there are no good structures available for hormone-sensitive lipase for example. Yet much is known about the structure and activity of lipases. Lipases have a catalytic Ser-His-Asp triad like serine proteases, which also carry out hydrolysis reactions. However, the substrates for lipase are nonpolar and could be part of larger structures such as lipid droplets. As such, the lipases work at the interface between aqueous and non-aqueous interfaces. They are interfacially activated. Lipases appear to have low activity in aqueous solutions but more in solutions that are more nonpolar. Many appear to have two conformations, a closed one with lower activity and poor lipid substrate binding properties, and an open one with higher activity and activation by nonpolar solvents. An alpha-helical lid appears to close off the active site in the closed conformation. In a more nonpolar solvent or the presents of bulk lipids, such as lipid droplets, the alpha-helical lid moves exposing the enzyme's hydrophobic binding site to substrate interactions. Figure \(17\) shows an interactive iCn3D model comparing the structure of a Thermomyces lanuginosus lipase in its closed (1DT3) and its open conformation (1EIN). Figure \(17\): The structure of a Thermomyces lanuginosus lipase in its closed (1DT3) and its open conformation (1EIN). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?C9yEwy6SR6MZ9Q1S7 The closed structure (1DT3) is shown in magenta and the open (1EIN) is in cyan. The active site triad Ser146, His258, and Asp201 is shown in CPK-colored sticks and labeled for each. The lid is shown in yellow with its side chains as sticks. Toggle between the two states using the "a" key. Note how the lid covers the active site in the closed (1DT3) structure. You will have to rotate the molecule to see the optimal orientation. Also, close out menu boxes for a larger and more centered display. Endocrine functions Not only does the adipocyte provide a store of energy but it is an endocrine organ that manages the flow of energy through the formation of the hormone leptin, which signals through the adiponectin receptors (AdipoR)1 and AdipoR2 and binds to the non-signaling interacting protein, T‑cadherin. The result is the stimulation of signaling cascades to communicate with other tissues through the secretion of cytokines and other mediators. The synthesis of leptin is tightly controlled by adipocytes mainly, although it is also produced by the stomach, placenta, and mammary gland, and its principal function is the provision of information on the state of fat stores to other tissues. In so doing, it regulates food intake and energy expenditure. Leptin was initially described as an anti-obesity hormone, but it serves as an adiposity signal with a vital function in maintaining adipose tissue mass to ensure survival under conditions of negative energy balance and so protect against either a deficit or an excess of adiposity. Lipid droplets have a role in this process, sinc perilipin is required for the sensing function. Adiponectin is a potent insulin sensitizer and suppressor of cell death and inflammation, directly promoting anti-diabetic and anti-atherosclerotic outcomes. Insulin is the main hormone that affects metabolism and its receptor at the plasma membrane is located in caveolae. The release of proinflammatory cytokines can stimulate lipolysis and cause insulin resistance, in turn leading to dysfunction of adipose tissue and systemic disruption of metabolism. Thus, adipose tissue metabolism has profound effects on whole-body metabolism, and defects in these processes can have severe implications for such serious pathological conditions as diabetes, obesity, cardiovascular disease, fatty liver disease, and cancer in humans. It is hoped that the development of specific inhibitors for hormone-sensitive lipase will improve the treatment of such metabolic complications. As caveolae, which contain the proteins caveolins (and presumably sphingolipids) and are particularly abundant in adipocytes, modulate the flux of fatty acids across the plasma membrane and are involved in signal transduction and membrane trafficking pathways, it is evident that they have a major role in this aspect of lipid metabolism. White fat acts as an endocrine organ and can release a variety of hormones including adipokines (analogous to cytokines, chemokines, lymphokines) adipsin, and leptin as well as tumor necrosis factor α (TNF-α), adiponectin, resistin, and RBP4. Functions other than energy management Lipid droplets accumulate within many cell types other than adipocytes, including leukocytes, epithelial cells, hepatocytes, and even astrocytes, especially during infections, cancer, and other inflammatory conditions. They are important for the cellular storage and release of hydrophobic vitamins, signaling precursors, and other lipids that are not related to energy homeostasis while reducing the dangers of lipotoxicity. On the other hand, excessive fatty acid accumulation is associated with lipotoxicity, endoplasmic reticulum stress, and mitochondrial damage and dysfunction, so lipid storage in lipid droplets must be balanced for health. A variety of enzymes are associated with lipid droplets, including protein kinases, which are involved in many different aspects of lipid metabolism, such as cell signaling, membrane trafficking, and control of the production of inflammatory mediators like the eicosanoids. Lipolysis enables the secretion of lipid species termed lipokines (more generally 'adipokines') from adipocytes that may signal in a hormone-like fashion to other tissues, thereby modulating gene expression and physiological function, including food intake, insulin sensitivity, insulin secretion, and related processes. These include palmitoleic acid (9-16:1) and fatty acid esters of hydroxy fatty acids (FAHFA), though the circulating proteins adiponectin and leptin have been studied more intensively. Adiponectin is a powerful insulin sensitizer and suppressor of apoptosis and inflammation with anti-diabetic and anti-atherosclerotic functions, often operating through its effects on sphingolipids, while leptin exerts most of its effects on the brain to trigger behavioral, metabolic, and endocrine responses to control the body's fuel reserves. Indeed, there are now suggestions that lipid droplets in all cell types are essential for the response mechanisms to cellular stress, including autophagy, inflammation, and immunity, and act as hubs to integrate metabolic and inflammatory processes. Via their lipolytic machinery, they regulate the availability of fatty acids for the activation of signaling pathways and the production of oxylipins from polyunsaturated fatty acids. For example, triacylglycerols in cytoplasmic lipid droplets of human mast cells, which are potent mediators of immune reactions and influence many inflammatory diseases, have a high content of arachidonic acid and this can be released by adipose triacylglycerol lipases as a substrate for production of specific eicosanoids when the cells are stimulated appropriately. Active metabolism in lipid droplets is important for the differentiation of monocytes, and it is essential for the sustained functional activity of differentiated macrophages, especially in relation to inflammation. During apoptosis, triacylglycerols enriched in polyunsaturated fatty acids accumulate in lipid droplets, possibly as a protective mechanism against membrane damage caused by oxidative stress and hydroperoxide formation in this process. Triacylglycerols in lipid droplets of the skin are a highly specific source of linoleic acid that is required for the formation of the O-acylceramides, which are essential for epidermal barrier function. An organelle termed the midbody in dividing cells in humans and rodents contains a unique triacylglycerol that is a single molecular species consisting of three fatty acids 16:1-12:0-18:1 (12:0 especially is rarely detected in human lipids), but its function is not known. Vitamin E (tocopherols) and vitamin A in the form of retinyl esters are stored in cytoplasmic lipid droplets, and the latter are present in appreciable concentrations in the stellate cells of the liver, for example. In endocrine cells of the gonads and adrenals, cholesterol esters stored in lipid droplets are an important source of cholesterol for the mitochondrial biosynthesis of various steroid hormones. In the nucleus of the cell, in addition to providing a reservoir of fatty acids for membrane remodeling, lipid droplets can sequester transcription factors and chromatin components and generate the lipid ligands for certain nuclear receptors. In addition to their role in lipid biochemistry, lipid droplets participate in protein degradation and glycosylation. Their metabolism can be manipulated by pathogenic viruses and bacteria such as Mycobacterium tuberculosis with unfortunate consequences for the host, but they also serve as reservoirs for proteins that fight intracellular pathogens. In consequence, such lipid droplets and their enzyme systems may be markers for disease states and are also considered to targets for pharmaceutical intervention. Insects In insects, the fat body is a multifunctional tissue that is the main metabolic organ. It integrates signals that control the immune system, molting, metamorphosis, and synthesis of hormones that regulate innumerable aspects of metabolism. In fat body cells, lipids, carbohydrates, and proteins are the substrates and products of many pathways for use in energy production or to act as reserves for mobilization at the appropriate stage of life (diapause, metamorphosis, and flight). In relation to innate and acquired humoral immunity, the fat body produces bactericidal proteins and polypeptides, i.e., lysozyme. It is also important in the early stages of an insect's life due to the production of vitellogenin, the yolk protein needed for the development of oocytes. Brown Adipose Tissue Most adipose tissue depots ('white fat') serve primarily as storage and endocrine organs that provide a reservoir of nutrients for release when the food supply is low. However, a second specialized form of adipose tissue, brown fat, is multilocular, highly vascularized, and rich in mitochondria and the iron-containing pigments that transport oxygen and give the tissue its color and name. Brown adipocytes arise from progenitor cells that are closer to those of skeletal muscle than white adipocytes. In humans, these depots tend to be located in specific anatomical regions such as subcutaneous areas around the neck, where their function may be to supply warm venous blood directly to the spinal cord and brain, and elsewhere to the heart, kidney, pancreas, and liver. Brown fat can oxidize fat so rapidly that heat is generated (“non-shivering thermogenesis”), and it is especially important in young animals and those recovering from hibernation. In brief, during cold exposure, the release of noradrenaline and stimulation of β-adrenergic receptors in the nervous system initiates a catabolic program that commences with a rapid breakdown of cellular triacylglycerol stores and release of unesterified fatty acids and transient activation of a co-activator of peroxisome proliferator-activated receptor gamma (PPARγ). These set in motion a signaling process that results in the efficient β-oxidation of fatty acids to produce heat. The key molecule is believed to be the uncoupling protein-1 (UCP1), which acts as a valve to uncouple electron transport in the respiratory chain from ATP production with a highly exothermic release of chemical energy, i.e., as heat rather than as ATP. Its actions are illustrated in Figure \(18\). Figure \(18\):  Role of UCP1 in generation of heat.  https://commons.wikimedia.org/wiki/F...n_the_cell.jpg.  Creative Commons Attribution-Share Alike 4.0 Internationa This is an example of a futile cycle that releases heat.  Although many aspects of the mechanism are uncertain, it is clear that proton conductance by UCP1 is highly regulated and inducible. It is activated by free long-chain fatty acids and inhibited by purine nucleotides, i.e., fatty acids are not only the substrate for thermogenesis but act also as self-regulating second messengers. The mitochondrial phospholipid cardiolipin, which is intimately involved in oxidative phosphorylation, is indispensable for stimulating and sustaining the function of thermogenic fat. Upon activation of brown adipose tissue, the dense vasculature increases the delivery of fatty acids and glucose to the brown adipocytes and warms the blood passing through the tissue. Long-chain fatty acids bind to UCP1 and may be transported through the membrane along with a proton.  Alternatively, it may just bind and activate the transport of a proton.  Figure \(19\) shows an interactive iCn3D model of the AlphaFold predicted structure of human UCP1 (P25874). Figure \(19\): AlphaFold predicted structure of human UCP1 (P25874). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...FkfrEqWE3D2dX6 Imagine a bilayer orienting perpendicularly to the gray helices representing the transmembrane helical segments that imbed the protein into the inner mitochondrial membrane.  The spacefill Ile150 denotes the matrix domain. Two key residues (K56, K29) involved in the binding of the carboxyl end of the fatty acids are shown on the matrix side of the protein is CPK-colored sticks and labeled. The black faces of the helices in the inner membrane region show the most probable binding site for the acyl chain of the fatty acid.  The lighter yellow/orange regions have greater predicted uncertainty in the AlphaFold computational model. There is evidence that acylcarnitines produced in the liver from fatty acids released from white adipose tissue in response to cold exposure are transported in plasma to brown adipose tissue and can serve as a substrate for thermogenesis. Indeed, a wide array of circulating lipids contributes to thermogenic potential, including free fatty acids and triacylglycerols. The activities of acyl-CoA synthetases and acyl-CoA thioesterases determine the availability of substrates for β-oxidation and consequently the thermogenic capacity. Synthesis of the lipokine (or 'batokine') octadecanoid 12,13-dihydroxy-9Z-octadecenoic acid (12,13-diHOME), is induced by cold also, and this stimulates the activity of brown adipose tissue by promoting the uptake of fatty acids, acting via G-protein-coupled receptors. It increases cardiac function and cardiomyocyte respiration via enhanced calcium cycling. Similarly, there are suggestions that n‑3 polyunsaturated fatty acids may promote adaptive thermogenesis, for example through the activity of the 12‑lipoxygenase metabolite and batokine 12‑hydroxyeicosapentaenoic acid (12-HEPE) by improving glucose metabolism via increased glucose uptake into adipocytes and skeletal muscle. (FAHFA) are relevant in this context. Peroxisomal synthesis of plasmenyl-phospholipids is believed to regulate adipose tissue thermogenesis by mediating mitochondrial fission. In hibernating mammals, brown adipose tissue is especially important metabolically, and even in laboratory animals such as mice, it can consume about 50% of dietary lipids and glucose when the animals are exposed to cold. Similarly, in even humans, the activity of brown adipose tissue is induced acutely by cold and is stimulated via the sympathetic nervous system, and the relevance of this tissue to human metabolism is now becoming apparent. For example, there are suggestions that brown adipose tissue can behave as an endocrine system to secrete endocrine factors ('batokines') that may be favorable in reducing cardiovascular risk. For obvious reasons, there are efforts to determine whether sustained activation of brown fat by pharmaceutical means could be beneficial for human disease states, including obesity, diabetes, and cardiovascular disease. Such research has been stimulated by the observation that clusters of distinct adipocytes with thermogenic capacity in addition to their storage function can be present in white adipose tissue and emerge in response to various physiological signals, especially reactive oxygen species. They are termed 'beige or brite' adipocytes and arise from multipotent pre-adipocytes. Brown adipose tissue is a discrete organ in animals, but beige adipose tissue is interspersed with white, and the two forms have different developmental origins. In adult humans, most of the deposits once thought to be classical brown adipose tissue do not contain the genetic markers for this tissue and are now recognized to be beige/brite fat, although brown adipose tissue per se is present in significant amounts in human newborns and infants. However, beige adipocytes utilize the same machinery to release heat by oxidation of fatty acids under β-adrenergic stimulation. The uncoupler protein 1 (UCP1) appears to be expressed only in brown and beige fat cells and is a marker for those cells.  It can be regulated transcriptionally and posttranscriptionally by ROS modification of a cysteine to produce a sulfenyl group.  The more classical brown adipocyte arises developmentally from the skeletal muscle-like cell, whereas the beige appears to develop to some extent from vascular smooth muscle cells. The beige cells can differentiate into more white adipocytes or more brown ones,  by the factors described in Figure \(20\). Figure (20\): Factors altering beige adipose tissue.  Paul Cohen and Bruce M. Spiegelman. Molecular Biology of the Cell, 27, 2017.  https://doi.org/10.1091/mbc.e15-10-0749. Attribution–Noncommercial–Share Alike 3.0 Unported Creative Commons License (http://creativecommons.org/licenses/by-nc-sa/3.0). Fat Cell Size In addition to the type of fat cell, the location and size of the adipocyte also affect health risks.  Obesity, cardiovascular disease, type II diabetes, and nonalcoholic fatty liver disease all depend on and are correlated with fat cell dysfunction.  The endocrine role of adipose tissue plays a key role in the development of dysfunction. The main depots of fat are the subcutaneous adipose tissue (SAT) and the visceral adipose tissue (VAT), the latter of which is associated with proinflammatory effects and associated greater health risks (type II diabetes, hypertension, metabolic syndrome, obesity, etc.).  Figure (21\) below summarizes the health consequence as the size of the SAT and VAT adipocytes increase (in the direction of the red arrow). Figure (2\): Adipocyte characteristics and cardiovascular risk. The diagram describes a proposed model, based on the results of adipocyte size/source, potential interactions with metabolic mediators, and pathophysiological effects. SAT, subcutaneous adipose tissue; VAT, visceral adipose tissue; HbA1c, glycated hemoglobin (a marker of elevated blood sugar associated with diabetes); RES, resistin; ADIPON, adiponectin; INS, insulin; HDLc; high-density lipoprotein cholesterol; FMD, flow-mediated dilation; CIMT, carotid intima media thickness.  Suárez-Cuenca et al. Sci Rep 11, 1831 (2021). https://doi.org/10.1038/s41598-021-81289-2.  Creative Commons Attribution 4.0 International License.   http://creativecommons.org/licenses/by/4.0/. It appears that the larger the adipocyte, the greater the inflammatory response. This condition is called adipocyte hypertrophy and it may be more important than obesity per se as a risk factor for inflammation, as indicated by inflammatory cytokine levels and macrophage infiltration into adipose tissue. The adipose-derived hormone leptin, when released under normal conditions, leads to appetite suppression and also decreases liver and muscle fat.  Higher fat storage is associated with increased blood leptin levels.   It also correlates with greater lymphocyte size. In contrast, the adipose hormone adiponectin leads to positive health consequences including increased sensitivity to insulin, decreased inflammation, and the formation of adipocytes from stem cells (adipogenesis).  The larger the fat cell, the lower the serum levels of adiponectin.  However, the link between fat cell size and adiponectin levels may be determined more by obesity. Other Functions of Triacylglycerol Depots Subcutaneous depots act as a cushion around joints and serve as insulation against the cold in many terrestrial animals, as is obvious in the pig, which is surrounded by a layer of fat, and it is especially true for marine mammals such as seals. Those adipocytes embedded in the skin differ from the general subcutaneous depots and support the growth of hair follicles and regenerating skin, and they may also have a defensive role both as a physical barrier and by responding metabolically to bacterial infection. In marine mammals and fish, the fat depots are less dense than water and so aid buoyancy with the result that less energy is expended in swimming. More surprisingly perhaps, triacylglycerols together with the structurally related glyceryl ether diesters and wax esters are the main components of the sonar lens used in echo location by dolphins and toothed whales. The triacylglycerols are distinctive in that they contain two molecules of 3‑methylbutyric (isovaleric) acid with one long-chain fatty acid. It appears that the relative concentrations of the various lipids in an organ in the head of the animals (termed the ‘melon’) are arranged anatomically in a three-dimensional topographical pattern to enable them to focus sound waves. In cold climates, many insects do not feed over winter and must manage their energy stores to meet the energetic demands of development and reproduction in the spring. Some insect species that are tolerant of freezing produce triacylglycerols containing acetic acid, and these remain liquid at low temperatures; by interacting with water, they may play a role in cryoprotection. Triacylglycerol Metabolism in Plants and Yeasts Fruit and seed oils are major agricultural products with appreciable economic and nutritional value to humans. The mesocarp of fruits is a highly nutritious energy source that attracts animals that help to disperse the seeds, and in plants such as the oil palm and olive trees a high proportion of the fruit flesh contains triacylglycerols. Similarly in seeds, triacylglycerols are the main storage lipid and can comprise as much as 60% of their weight. Fruit lipids are not intended for use by the plant per se and are stored in lipid droplets in large irregular structures that break down readily, but seed lipids are required for the development of the plant embryo, so their metabolism is of particular importance. However, triacylglycerol biosynthesis and metabolism are required also for pollen viability and to maintain lipid homeostasis in chloroplasts (see the note on plastoglobules below). Seed oils: In seeds and other plant tissues, biosynthesis of fatty acids takes place in plastids, and these are stored in the form of triacylglycerols in lipid droplets with a coherent surface layer of proteins and lipids in the embryo (e.g., Arabidopsis, soybean or sunflower) or endosperm (e.g., castor bean) tissues of seeds. In addition to the common range of fatty acids synthesized in plastids, mainly palmitate, and oleate, some plant species produce novel fatty acids, including medium- and very-long-chain components and those with oxygenated and other functional moieties. A specific means of diverting these to seeds for triacylglycerol production exists to prevent disruption of the plant membranes. Seed development occurs in three stages - rapid cell division with no accumulation of storage material, rapid deposition of triacylglycerols and other energy-rich metabolites, and finally desiccation. During the period of oil accumulation in seeds, the newly formed ACP esters of fatty acids are first hydrolyzed by two different classes of acyl-ACP thioesterases at the inner plastid envelope membrane, before the unesterified fatty acids are transported to the endoplasmic reticulum (ER), by a family of fatty acid export proteins (FAX) of which there are seven isoforms in Arabidopsis, two of which (FAX2 and FAX4) are highly expressed during the early stage of seed development. The unesterified fatty acids are shuttled across the plastid outer envelope, probably by vectorial acylation by long-chain acyl-CoA synthases, which catalyze the formation of CoA esters. In the ER, triacylglycerols and membrane lipids are synthesized by the Kennedy and other pathways described above. In yeast and plants, 1,2‑diacyl-sn-glycerol esterification is the only committed step in triacylglycerol production, and this occurs by mechanisms that can be both dependent or independent of acyl-CoA esters. The acyl-CoA-dependent route is catalyzed by diacylglycerol:acyl-CoA acyltransferases (DGATs) with acyl-CoA and diacylglycerols as substrates, and two membrane-bound isoenzymes (DGAT1 and DGAT2) and a cytosolic isoenzyme (DGAT3) are known, DGAT1 is a key enzyme involved in triacylglycerol formation in developing seeds, while DGAT2 is especially important in those plant species with unusual fatty acid compositions. In Arabidopsis, DGAT3 has some specificity for polyunsaturated fatty acids in seed development. Figure \(22\) summarizes TAG synthesis in plant ER. In addition, a substantial proportion of triacylglycerol biosynthesis in some plant species is synthesized by flux through the membrane phospholipid phosphatidylcholine, produced by what are sometimes termed inaccurately the 'eukaryotic and prokaryotic pathways' with differing positional distributions, in which diacylglycerols are generated from phosphatidic acid by the action of a phosphatidate phosphatase as an intermediate. In the acyl-CoA-independent reaction, the direct transfer of one fatty acid from phosphatidylcholine to diacylglycerol by the action of the phospholipid:1,2-diacyl-sn-glycerol-acyltransferase (PDAT) enzyme also occurs, with the formation of lysophosphatidylcholine as a byproduct, which can be re-esterified for further reaction. As phosphatidylcholine undergoes extensive remodeling and its fatty acid components are subject to modification, for example by desaturation to form linoleic and linolenic acids, the compositions and especially the positional distributions of triacylglycerols produced in this way can be very different from those synthesized by the ‘classical’ pathways. Phosphatidylcholine may also function as a carrier for the trafficking of acyl groups between organelles and membrane subdomains, and it has been suggested that an assembly of interacting enzymes may facilitate the transfer of polyunsaturated fatty acids from this phospholipid to triacylglycerols in seeds. Both sterol and sphingolipid biosynthesis appear to be important factors for efficient seed oil production. As triacylglycerol synthesis continues, oil droplets accumulate between the leaflets of the endoplasmic reticulum and are surrounded by a monolayer of phospholipids, sterols, and proteins, which in Arabidopsis include oleosins, a caleosin, a steroleosin, a putative aquaporin, and a glycosylphosphatidylinositol-anchored protein. Oleosins are the most abundant of these (~65%) and are small proteins (15-30 kDa) that contain cytosolic-facing N- and C-termini and a large hydrophobic domain necessary to target them to lipid droplets, where they are important for control of their size and stability. Eventually, lipid droplets “bud off” from the endoplasmic reticulum with their monolayer of phospholipids and proteins, and they are released into the cytosol by a yet-to-be-defined mechanism. At the onset of germination, water is absorbed, and esterases/lipases are activated. The process of lipolysis begins at the surface of oil bodies, where the oleosins, which are the most abundant structural proteins, are believed to assist in the docking of lipases and to control the size and stability of lipid droplets in seeds. A number of esterases/lipases have been cloned from various plant species and possess a conserved catalytic triad of Ser, His, and Asp or Glu, somewhat different from the animal lipases, as in patatin (an especially abundant lipolytic protein in potatoes), which can hydrolyze triacylglycerols but not phospho- or galactolipids. The most important of these is believed to be the 'sugar-dependent lipase 1 (SDP1)', which is a patatin-like lipase similar in function to the mammalian adipose triacylglycerol lipase discussed above and is located on the surface of the oil body. This is active mainly against triacylglycerols to generate diacylglycerols but presumably works in conjunction with di- and monoacylglycerol lipases to generate free fatty acids and glycerol. The lipid droplets in seeds exist near glyoxysomes (broadly equivalent to peroxisomes). These are the membrane-bound organelles that contain most of the enzymes required to oxidize fatty acids derived from the triacylglycerols via acetyl-CoA to four-carbon compounds, such as succinate, which are then converted to soluble sugars to provide germinating seeds with energy to fuel the growth of the seedlings and to produce shoots and leaves. In addition, they supply structural elements before the seedlings develop the capacity to photosynthesize. How the products of lipolysis are transported to the glyoxysomes for further metabolism has still to be determined, but a specific ‘ABC’ transporter is required to import fatty acids into the glyoxysomes in Arabidopsis. The free acids are converted to their coenzyme A esters by two long-chain acyl-CoA synthetases located on the inner face of the peroxisome membrane before entry into the β‑oxidation pathway. All these processes are controlled by an intricate regulatory network, involving transcription factors that crosstalk with signaling events from the seed maturation phase through to embryo development. After about two days of the germination process, the glyoxysomes begin to break down, but β-oxidation can continue in peroxisomes in leaf tissue. Lipid droplets - plastoglobules: Triacylglycerol-rich lipid droplets (LD) have been observed in most cell types in vegetative tissues of plants as well as in seeds, and although their origin and function are poorly understood, they contain all the enzymes required for triacylglycerol metabolism together with phospholipases, lipoxygenases, and other oxidative enzymes. Instead of oleosins, these lipid droplets in plants and algae contain a family of ubiquitously expressed 'LD-associated proteins' on the surface, together with a monolayer of phospholipids (mainly phosphatidylcholine), galactolipids such as sulfoquinovosyldiacylglycerol and in some species betaine lipids. As in yeast and humans, seipins (three in Arabidopsis) are necessary for normal LD biogenesis. Again, LD forms within the bilayer of the endoplasmic reticulum and pinch off into the cytoplasm. Abiotic stresses can induce the remodeling of lipid membranes through lipase action with the formation of toxic lipid intermediates, and these can be sequestered by triacylglycerols in lipid droplets to inhibit membrane damage and potentially prevent cell death. While they are believed to be involved mainly in stress responses, lipid droplets may have other specialized roles, for example in anther and pollen development, where triacylglycerols serve as a source of fatty acids for membrane biosynthesis. Fatty acids derived from triacylglycerols in lipid droplets are believed to be subjected to peroxisomal β‑oxidation to produce the ATP required for stomatal opening and no doubt many other purposes. In addition, lipid droplets that have been termed 'plastoglobules' are produced by a localized accumulation of triacylglycerols and other neutral lipids between the membrane leaflets of the thylakoid cisternae and then pinch off into the stroma, where they are involved in a wide range of biological functions from biogenesis to senescence via the recruitment of specific proteins. During senescence, for example, lipid droplets accumulate rapidly in the leaves of A. thaliana. In reproductive tissues, may have a more direct function by recruiting and transporting proteins, both for organ formation and successful pollination. Antifungal compounds such as 2-hydroxy-octadecatrienoic acid and other oxylipins are produced from α-linolenic acid in these organelles, and it has been suggested that the latter function as intracellular factories to produce stable metabolites via unstable intermediates by concentrating the enzymes and hydrophobic substrates efficiently. Plastoglobules are also implicated in the biosynthesis and metabolism of vitamins E and K. Microalgae: Triacylglycerol metabolism in lipid droplets in microalgae is under intensive study because of their potential for nutraceutical and biodiesel production. It seems that similar processes occur in higher plants, but with a simpler genome encoding few redundant proteins. In the unicellular green model microalga Chlamydomonas reinhardtii, for example, key lipid droplet proteins, lipases, and enzymes of β-oxidation have been characterized. Yeasts: Lipid droplets in yeast are a highly dynamic and functionally diverse hub that ensures stress resistance and cell survival by promoting membrane and organelle homeostasis. As most of the important biosynthetic and catabolic enzymes involved in triacylglycerol metabolism are conserved between yeasts and mammals, the former proving to be useful models for the study of triacylglycerol production. The size and triacylglycerol content of lipid droplets in yeasts change appreciably in different stages of growth and development, and Saccharomyces cerevisiae contains a single phosphatidic acid phosphatase (Pah1), which has an essential role in this process. During vegetative growth, Pah1 in the cytosol is phosphorylated by multiple protein kinases, and this enables the synthesis of phospholipids rather than triacylglycerols. As cells progress into stasis, the Pah1 is dephosphorylated and translocates to the endoplasmic reticulum, which ultimately leads to triacylglycerol synthesis for storage in lipid droplets. Some fatty acids derived from phospholipids are utilized for triacylglycerol biosynthesis at the inner nuclear membrane, and this is important for nuclear integrity. Triacylglycerol Metabolism in Prokaryotes The study of the biosynthesis of triacylglycerols in bacteria has been stimulated by an awareness of the role of this lipid class in the pathogenesis of Mycobacterium tuberculosis and the relationship with antibiotic biosynthesis by Streptomyces coelicolor. For example, triacylglycerols are believed to be an energy reserve for the long-term survival of M. tuberculosis during the persistence phase of infection as well as a means by which unesterified fatty acids are detoxified. Increasing numbers of bacterial species, for example from the genera Mycobacterium, Nocardia, Rhodococcus, Micromonospora, Dietzia, and Gordonia, are now known to produce triacylglycerols (sometimes wax esters), and these can be stored in lipid droplets in the organisms. The first three steps in triacylglycerol biosynthesis are catalyzed by GPAT, LPAT, and PAP enzymes comparable to those in other organisms. However, it has become apparent that the DGAT can be a dual-function CoA-dependent acyltransferase known as wax ester synthase/diacylglycerol acyltransferase, which accepts a broad diversity of acyl-CoA substrates for esterification of diacylglycerols or long-chain fatty alcohols for the synthesis of triacylglycerols or wax esters, respectively, depending on which intermediates are present in the organisms. Bacteria that lack such an enzyme are unable to produce these non-polar lipids.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/21%3A_Lipid_Biosynthesis/21.02%3A_Biosynthesis_of_Triacylglycerols.txt

Search Fundamentals of Biochemistry by William (Bill) W. Christie and Henry Jakubowski. This section is an abbreviated and modified version of material from the Lipid Web, an introduction to the chemistry and biochemistry of individual lipid classes, written by William Christie. Introduction In this section will be explore the synthesis of the membrane glycerophospholipids and their metabolic derivatives. We will start with the simplest one, phosphatidic acids, and end with phosphatidylinositols. Phosphatidic Acids and Derivatives Phosphatidic acid or 1,2-diacyl-sn-glycero-3-phosphate is a key intermediate in the biosynthesis both of other glycerophospholipids and of triacylglycerols. It is structurally one of the simplest of the phospholipids and was long thought to be important only as a precursor of other lipids, where it is indeed a key molecule, but it is now known to have many other functions in animals, plants, and other organisms by its influence on membrane structure and dynamics, and by its interactions with various proteins. As a lipid mediator, it modulates various signaling and cellular processes, such as membrane tethering, conformational changes and enzymatic activities of specific proteins, and vesicular trafficking. Moreover, its metabolite lysophosphatidic acid is recognized as a key signaling molecule with a myriad of biological effects mediated through specific receptors. Phosphatidic Acid – Occurrence and Biosynthesis Phosphatidic acid is not an abundant lipid constituent of any living organism, seldom greater than picomolar concentrations in cells, but it is extremely important both as an intermediate in the biosynthesis of other glycerophospholipids and triacylglycerols and as a signaling molecule or a precursor of signaling molecules. Indeed, it is often over-estimated in tissues as it can arise by inadvertent enzymatic hydrolysis during inappropriate storage or extraction conditions during analysis. It is the simplest diacyl-glycerophospholipid, and the only one with a phosphomonoester as the head group. The molecule is acidic and carries a negative charge, i.e., it is an anionic lipid. The structure of phosphatidic acid is shown in Figure \(1\). There are at least four important biosynthetic pathways for phosphatidic acid biosynthesis in different organelles under various stimuli, and possibly resulting in the formation of different molecular species. The main pathway involves sequential acylation of sn-glycerol-3-phosphate, derived from catabolism of glucose, by acyl-coA derivatives of fatty acids. First, one acyltransferases catalyses the acylation of position sn-1 to form lysophosphatidic acid (1‑acyl-sn-glycerol-3-phosphate), and then a second specific acyltransferase catalyses the acylation of position sn-2 to yield phosphatidic acid. The synthesis of phosphatidic acid from glycerol-3-phosphate is shown in Figure \(2\). In mammals, the glycerol-3-phosphate acyltransferase that catalyses the first step exists in four isoforms, two in the mitochondrial outer membrane (designated GPAT1 and 2) and two in the endoplasmic reticulum (GPAT3 and 4); all are membrane-bound enzymes, which are believed to span the membranes. GPAT1 is highly expressed in the liver and adipose tissue, where it is responsive to changes in feeding status via the sterol regulatory element binding protein-1 (SREBP-1), a master transcriptional regulator of lipogenic enzymes. It is essential in directing fatty acyl-CoA esters towards glycerolipid synthesis as opposed to β-oxidation. GPAT3 is especially important for triacylglycerol storage in adipocytes, while GPAT4 is the main contributor to lysophosphatidic acid synthesis in liver and brown adipose tissue. For the second step in phosphatidic acid biosynthesis, five mammalian acyl-CoA:lysophosphatidic acid acyltransferases are known of which three are in the endoplasmic reticulum (LPAAT or LPAT or AGPAT1, 2 and 3), with a further two (LPAT4 and 5) on the outer mitochondrial membrane. While LPAT1 and 2 have strict specificity for lysophosphatidic acid as acyl acceptor, other isoforms can esterify other lysophospholipids. Human LPAT1 showed higher activity with 14:0-, 16:0- and 18:2‑CoAs, while LPAT2 prefers 20:4-CoA and LPAT3 produces phosphatidic acid containing docosahexaenoic acid (22:6(n-3)); the last is especially important in retina and testes. LPAT4 and 5 have a preference for oleoyl-CoA and polyunsaturated acyl-CoAs as the acyl donor, suggesting a dual role in glycerolipid synthesis and remodeling. The activity in the endoplasmic reticulum predominates in adipose tissue, but the mitochondrial forms are believed to be responsible for half the activity in liver. However, as there is traffic of phosphatidic acid between the mitochondria and endoplasmic reticulum for remodeling or for synthesis of other lipids, the relative contributions of the two can be difficult to assess. In plants, the sn-glycerol-3-phosphate pathway exists both in plastids and at the endoplasmic reticulum with multiple isoforms of the two acyltransferases as well as differences in the acyl substrates. In brief most plant lipid biosynthesis begins with fatty acid biosynthesis in the chloroplasts. In plastids, the acyltransferase ATS1 transfers 18:1 acyl groups from acyl-acyl carrier protein (acyl-ACP) to position sn-1 of glycerol 3-phosphate, before ATS2 transfers a palmitoyl group from ACP to position sn-2, producing phosphatidic acid at the inner leaflet of the chloroplast inner envelope membrane (IEM). Fatty acids intended for the endoplasmic reticulum are released from ACP in the chloroplast stroma by IEM-associated thioesterases, exported and then activated by acyl-CoA synthetases of the outer envelope membrane to produce species with C18 fatty acids in both positions. Thus, acyl-CoAs are used for phosphatidic acid biosynthesis in the endoplasmic reticulum with marked differences in the specificity of the acyl substrates. Subsequently, phosphatidic acid in the plastids is utilized for biosynthesis of galactosyldiacylglycerols, while that in the endoplasmic reticulum is used for synthesis of triacylglycerols and phospholipids. In bacteria, two families of enzymes are responsible for acylation of position sn-1 of glycerol-3-phosphate. One present in Escherichia coli, for example, utilizes the acyl-acyl carrier protein (acyl-ACP) products of fatty acid synthesis as acyl donors as well as acyl-CoA derived from exogenous fatty acids. In a second wider group of bacteria, including cyanobacteria, there are enzymes (PlsX and PlsY) that make use of the unique acyl donors, acyl-phosphates derived in part from acyl-ACP, to acylate position sn-1. Acylation of position sn-2 in this instance is performed by a further family of enzymes (PlsC) that uses acyl-ACP as the acyl donor, although some bacterial species may use acyl-CoA also. In animals, a second biosynthetic pathway utilizes dihydroxyacetone phosphate (DHAP) as the primary precursor for the peroxisomal enzyme, DHAP acyltransferase, which produces acyl-DHAP. This intermediate is converted to lysophosphatidic acid in a NADPH-dependent reaction catalyzed by acyl-DHAP reductase, and this is in turn acylated to form phosphatidic acid by the same LPAT as in the previous mechanism. This pathway is of particular importance in the biosynthesis of ether lipids. The synthesis of phosphatidic acid from dihydroxyacetone phosphate is shown in Figure \(3\). A third important route to phosphatidic acid is via hydrolysis of other phospholipids, but especially phosphatidylcholine, by the enzyme phospholipase D (or by a family or related enzymes of this kind). The enzyme is readily available for study in plants, where the special functions of phosphatidic acid have long been known (see below), but it is now recognized that phospholipase D is present in bacteria, yeasts and most animal cells. In the last, it exists in two main isoforms with differing specificities and cellular locations; PLD1 is found mainly in the Golgi-lysosome continuum, while PLD2 is present mainly in the plasma membrane. They are phosphoproteins, the activity of which is regulated by kinases and phosphatases and by binding to phosphatidylinositol-4,5-bisphosphate. In mitochondria, a distinctive enzyme of this type utilizes cardiolipin as substrate. The mechanism involves the use of water as the nucleophile to catalyse the hydrolysis of phosphodiester bonds in phospholipids. Phospholipase D activity is dependent on and regulated by neurotransmitters, hormones, small monomeric GTPases and lipids. The hydrolysis of phosphatidylcholine to phosphatidic acids by phospholipase D is shown in Figure \(4\). In addition to its function in generating phosphatidic acid mainly for signaling purposes but also for the maintenance of membrane composition, phospholipase D is involved in intracellular protein trafficking, cytoskeletal dynamics, cell migration, and cell proliferation, partly through protein-protein interactions; it is considered to be important in inflammation and in cancer growth and metastasis as a downstream transcriptional target of proteins involved in the pathophysiology of these diseases. It also has an unusual activity as a guanine nucleotide exchange factor. By a transphosphatidylation reaction with ethanol, it generates phosphatidylethanol, a useful biomarker for ethanol consumption in humans. Under some conditions, phosphatidic acid can be generated from 1,2-diacyl-sn-glycerols by the action of diacylglycerol kinases, for example those produced from other phospholipids by the action of phospholipase C. Such enzymes appear to be ubiquitous in nature, although those in bacteria and yeast are structurally different from the mammalian enzymes. Diacylglycerol kinases, of which at least ten isoforms (DGKα to DGKκ) exist with different sub-cellular locations and functions in animals, use ATP as the phosphate donor. While the epsilon isoform (DGKε) utilizes the 1-stearoyl-2-arachidonoyl species of diacyl-sn-glycerols preferentially to produce phosphatidic acid for the biosynthesis of phosphatidylinositol, other isoenzymes phosphorylate diverse diacylglycerol species. Aside from producing phosphatidic acid for phospholipid production or signaling , these enzymes may attenuate the signaling effects of diacylglycerols. For example, diacylglycerol kinases can contribute to cellular asymmetry and control the polarity of cells by regulating the gradients in diacylglycerol and phosphatidic acid concentrations. Figure \(5\) shows the synthesis of phosphatidic acid via diacylglycerols and the reverse reaction. The reverse reaction, hydrolysis, is catalyzed by lipins. These enzymes are of importance in regulating the local concentrations of phosphatidic acid and thence its biological activity. A further possible route to phosphatidic acid production for signaling specifically is via acylation of lysophosphatidic acid, which can be produced independently for signaling purposes as discussed below. This pathway may be especially relevant in membranes, where the protein endophilin has LPAT activity and is believed to generate phosphatidic acid from lysophosphatidic acid in order to alter the curvature of the membrane bilayer. Phosphatidic Acid - Role as a Lipid Precursor In summary, phosphatidic acid generated via 1-acyl-sn-glycerol-3-phosphate is the primary precursor of other glycerolipids, although other pathways may be more important for generating the lipid for signaling functions. Whether separate pools of this lipid for specific purposes really exist is not certain since dynamic changes of intracellular distribution occur under various cellular conditions. These are attributed to inter-organelle transfer via vesicular transport or at membrane contact sites by lipid transfer proteins. Control of its concentration in membranes, especially in the endoplasmic reticulum, is therefore of great importance, and a transcriptional repressor 'Opi1', which binds specifically to phosphatidic acid in membranes, is a key regulatory factor. However, many other phosphatidic acid-binding proteins have been identified that influence how phosphatidic acid is used either as a biosynthetic precursor or for signaling purposes. The mechanisms for phosphatidic acid homeostasis differ among animals, plants, yeasts, and bacteria in response to the differing functional requirements in these organisms. Figure \(6\) shows the pathways for biosynthesis of complex glycerolipids. In addition to dietary, hormonal and tissue-specific factors in animals, the extent to which fatty acids are channeling either into triacylglycerol synthesis for storage in lipid droplets and secretion in lipoproteins or into glycerophospholipids for membrane formation depends to a large extent upon the enzymes of glycerol-3-phosphate pathway, their isoform expression, activities and locations. On the other hand, phosphatidic acid is not only a biosynthetic precursor of other lipids but also a regulatory molecule in the transcriptional control of the genes for glycerolipid synthesis, and regulation of its concentration in cells for this purpose is similarly essential. For example, the local concentration of phosphatidic acid in the endoplasmic reticulum is an important factor in the biogenesis of lipid droplets. The subsequent steps in the utilization of phosphatidic acid in the biosynthesis of triacylglycerols and of the various glycerophospholipids are described in separate documents of this website. Thus, hydrolysis of phosphatidic acid by phosphatidate phosphatase enzymes (including lipins 1, 2, and 3) is the source of most other glycerolipids, e.g. sn‑1,2‑diacylglycerols (DG), which are the precursors for the biosynthesis of triacylglycerols (TAG), phosphatidylcholine (PC) and phosphatidylethanolamine (PE) via the so-called Kennedy pathway (also of monogalactosyldiacylglycerols in plants). Via reaction with cytidine triphosphate, phosphatidic acid is the precursor of cytidine diphosphate diacylglycerol, which is the key intermediate in the synthesis of phosphatidylglycerol (PG), and thence of cardiolipin (CL), and of phosphatidylinositol (PI), and in prokaryotes and yeast but not animals phosphatidylserine (PS). Depending on the organism and other factors, phosphatidylserine can be a precursor for phosphatidylethanolamine, while the latter can give rise to phosphatidylcholine by way of mono- and dimethyl-phosphatidylethanolamine intermediates. The cytidine diphosphate diacylglycerol synthase is another enzyme that consumes phosphatidic acid and is important for modulating the concentration of phosphatidic acid in cells and for regulating processes mediated by this lipid. While the fatty acid composition of phosphatidic acid can resemble that of the eventual products, the latter are generally much altered by re-modeling after synthesis via deacylation-reacylation reactions. Phosphatidic Acid - Biological Functions in Animals In addition to its role as an intermediate in lipid biosynthesis, phosphatidic acid and especially that generated by the action of phospholipase D and by diacylglycerol kinases may have signaling functions as a second messenger, although it is not certain whether all the activities suggested by studies in vitro operate in vivo. Nonetheless, phosphatidic acid has been implicated in many aspects of animal cell biochemistry and physiology. Some of the observed effects may be explained simply by the physical properties of phosphatidic acid, which has a propensity to form a hexagonal II phase, especially in the presence of calcium ions. Thus, hydrolysis of phosphatidylcholine, a cylindrical non-fusogenic lipid, converts it into cone-shaped phosphatidic acid, which promotes negative membrane curvature and fusion of membranes. It differs from other anionic phospholipids in that its small anionic phosphomonoester head group lies very close to the hydrophobic interior of the lipid bilayer. In model systems, phosphatidic acid can effect membrane fusion, probably because of its ability to form non-bilayer phases. For example, the phosphatidic acid biosynthesis is believed to favor intraluminal budding of endosomal membranes with the formation of exosomes, and in many cell types, vesicle trafficking, secretion and endocytosis may require phosphatidic acid derived by the action of phospholipase D. Also of relevance in this context is its overall negative charge, and it is not always clear whether some of the observed biological effects are specific to phosphatidic acid or simply to negatively charged phospholipids in general. In contrast to phosphoinositide-interacting proteins, which have defined structural folds, the binding motifs of effector proteins with phosphatidic acid are not highly conserved. However, it has been demonstrated that the positively charged lysine and arginine residues on proteins can bind with some specificity to phosphatidic acid through hydrogen bonding with the phosphate group thus distinguishing it from other phospholipids. An ‘electrostatic-hydrogen bond switch model’ has been proposed in which the head group of phosphatidic acid forms a hydrogen bond to amino acid residues leading to de-protonation of the head group, increasing its negative charge from -1 to -2 and thus enabling stronger interactions with basic residues and tight docking with the membrane interacting protein. In this way, phosphatidic acid can tether certain proteins to membranes, and it can simultaneously induce conformational changes, hinder ligand binding and/or oligomerize proteins to alter their catalytic activity, stability and interactions with other molecules. It functions as a cellular pH sensor in effect in that binding to proteins is dependent on intracellular pH and the protonation state of its phosphate head group. One key target of the lipid is mTOR, a serine/threonine protein kinase with a signaling cascade that regulates cell growth, proliferation, motility and survival, together with protein synthesis and transcription, by integrating both nutrient and growth factor signals. This forms two distinct complexes of accessory proteins that regulate downstream targets. Of these, mTORC1 interacts directly with phosphatidic acid and this interaction allosterically activates the enzyme complex to regulate protein synthesis, mitochondrial metabolism and the transcription of enzymes involved in lipid synthesis. In contrast, phosphatidic acid appears to inhibit mTORC2 activity, for example in relation to insulin signaling . Phosphatidic acid is believed to regulate membrane trafficking events, and it is involved in activation of the enzyme NADPH oxidase, which operates as part of the defence mechanism against infection and tissue damage during inflammation. By binding to targeted proteins, including protein kinases, protein phosphatases and G-proteins, it may increase or inhibit their activities. Effects on gene transcription have been observed that are linked to inhibition of peroxisome proliferator-activated receptor (PPAR) activity. In yeast, phosphatidic acid in the endoplasmic reticulum binds directly to a specific transcriptional repressor to keep it inactive outside the nucleus; when the lipid precursor inositol is added, this phosphatidic acid is rapidly depleted, releasing the transcriptional factor so that it can be translocated to the nucleus where it is able to repress target genes. The overall effect is a mechanism to control phospholipid synthesis. In addition, phosphatidic acid regulates many aspects of phosphoinositide function. For example, the murine phosphatidylinositol 4-phosphate 5-kinase, the main enzyme generating the lipid second messenger phosphatidylinositol-4,5-bisphosphate, does not appear to function unless phosphatidic acid is bound to it; this lipid, generated by the action of phospholipase D, recruits the enzyme to the membrane and induces a conformational change that regulates its activity. It may have a role in promoting phospholipase A2 activity, a key enzyme in eicosanoid production from phosphoinositide precursors. In relation to signaling activities, it should be noted that phosphatidic acid can be metabolized to sn-1,2-diacylglycerols or to lysophosphatidic acid (see next section), both of which have distinctive signaling functions in their own right. Conversely, both of these compounds can be in effect be de-activated by conversion back to phosphatidic acid. Phospholipase D isoforms and phosphatidic acid have been implicated in a variety of pathologies including neurodegenerative diseases, blood disorders, late-onset Alzheimer's disease and cancer, leading to attempts to develop specific inhibitors of the enzyme for therapeutic purposes. Similarly, the expression of LPAT isoforms can enhance the proliferation and chemoresistance of some cancer cells. Diacylglycerol kinase alpha (DGKα) is highly expressed in several refractory cancer cells, where it attenuates apoptosis, and promotes proliferation. In addition, DGKα is highly abundant in T cells and induces a nonresponsive state, which enables advanced cancers to escape immune action. Inhibition of this enzyme also is seen as a promising treatment strategy. Phosphatidic Acid - Biological Functions in Plants Phosphatidic acid is present at higher levels in roots of plants in comparison to leaves and is believed to have a function in root architecture. Similarly, its concentration is elevated in flowers and reproductive tissues, but the significance of this is not known. In addition to its role as one of the central molecules in lipid biosynthesis, it facilitates the transport of lipids across plant membranes, and it is also the key plant lipid second messenger, which is rapidly and transiently generated in response to many different biotic and abiotic stresses. In contrast to animal metabolism, the diacylglycerol signaling pathway is believed to be relatively insignificant in plants. The main source of phosphatidic acid for these purposes is the action of phospholipase D (PLD) on membrane phospholipids, such as phosphatidylcholine and phosphatidylethanolamine. Plants contain numerous related enzymes of this type, 12 in Arabidopsis and 17 in rice, in comparison with two in humans and one in yeast, and individual iso-enzymes may elicit specific responses. In the former, the isoforms are grouped into six classes, based on the genic architecture, sequence similarities, domain structures and biochemical properties. These depend mainly on their lipid-binding domains, with some homologous to the human and yeast enzymes and with most containing a characteristic ‘C2’ (calcium- and lipid-binding) domain. The most widespread of these is PLDα, which does not require binding to phosphatidylinositol 4,5-bisphosphate, in contrast to other PLD isoforms and the mammalian enzyme, but millimolar levels of Ca2+ are necessary. Studies with fluorescent biosensors suggest that phosphatidic acid accumulates in the subapical region of the cytosolic leaflet of the plasma membrane. Phosphatidic acid can also be produced by the sequential action of phospholipase C and diacylglycerol kinase on membrane inositol phospholipids, with diacylglycerols as an intermediate (there are 7 isoenzymes in A. thaliana). One difference from animal metabolism is that diacylglycerol pyrophosphate can be synthesized from phosphatidic acid in plants (see below). Phosphatidic acid is required to bind and allosteric activate the monogalactosyldiacylglycerol synthase (MGDG1), located in the inner envelope membrane of the chloroplast, and it may be a regulator of the biosynthesis of thylakoid membranes. Phospholipase D activity and the phosphatidic acid produced have long been recognized as of importance during germination and senescence, and they have an essential role in the response to stress damage and pathogen attack, both in higher plants and in green algae. A high content of phosphatidic acid induced by phospholipase D action during wounding or senescence brings about a loss of the membrane bilayer phase, because of the conical shape of this negatively charged phospholipid in comparison to the cylindrical shape of structural phospholipids. This change in ionization properties has crucial effects upon lipid-protein interactions, "the electrostatic-hydrogen bond switch model" described above. By promoting negative curvature at the plasma membrane and binding to clathrin proteins, it is believed to facilitate the process of endocytosis. Similar phenomena may explain why phosphatidic acid is important in the response to other forms of stress, including osmotic stress (salinity or drought), cold and oxidation. Although much remains to be learned of the mechanism by which it exerts its effects, it is believed to promote the response to the plant hormone abscisic acid. In addition, phosphatidic acid may interact with salicylic acid to mediate defence responses. In plants, phosphatidic acid is involved in many different cell responses induced by hormones, stress and developmental processes. In relation to cellular signaling, it often acts in concert with phosphatidylinositol 4,5-bisphosphate by binding to specific proteins rather than acting via a receptor. As in mammalian cells, targets for such signaling include protein kinases and phosphatases in addition to proteins involved in membrane trafficking and the organization of the cytoskeleton. It can both activate or inhibit enzymes. If the target protein is soluble, binding to phosphatidic acid can cause the protein to be sequestered into a membrane with effects upon downstream targets. For example, it is involved in promoting the growth of pollen-tubes and root hairs, decreasing peroxide-induced cell death, and mediating the signaling processes that lead to responses to ethylene and again to the hormone abscisic acid. Thus, in the 'model' plant Arabidopsis, phosphatidic acid interacts with a protein phosphatase to signal the closure of stomata promoted by abscisic acid; it interacts also with a further enzyme to mediate the inhibition of stomatal opening effected by abscisic acid. Together these reactions constitute a signaling pathway that regulates water loss from plants. It is noteworthy that phosphatidic acid production can be initiated by opposing stress factors, such as cold and heat, as well as by hormones that are considered to be antagonistic, such as abscisic acid and salicylic acid. It is possible that phosphatidic acid molecules synthesized by the two main pathways differ in composition and cellular distributions and so may produce different responses, but this is an open question. Certainly, during low temperature stress, phosphatidic acid is generated by the action of diacylglycerol kinase. It also seems likely that these differing activities are controlled by the cellular environment where the lipid is produced and by the availability of target proteins or other molecules with which it can act synergistically. Genes encoding enzymes involved in phosphatidic acid metabolism have been manipulated to explore their potential application for crop improvements, based on effects on plant growth, development, and stress responses. As in animals, phosphatidic acid is catabolized and its signaling functions are terminated by lipid phosphate phosphatases and phosphatidic acid hydrolases, and by acyl-hydrolases and lipoxygenases with the production of fatty acids and other small molecules, which are subsequently absorbed and recycled. Lysophosphatidic Acid Figure \(7\) shows the structure of a lysophosphatidic acid (note the absence of an acyl group at C2). Lysophosphatidic acid (LPA) or 1-acyl-sn-glycerol-3-phosphate differs structurally from phosphatidic acid in having only one mole of fatty acid per mole of lipid. As such, it is one of the simplest possible glycerophospholipids. It exists in the form of many different molecular species, i.e., esterified to 16:0 to 22:6 fatty acids, and there is preliminary evidence that saturated and polyunsaturated species may differ in their biological properties in some circumstances. As the sn-1-acylated form is more stable thermodynamically, facile isomerization ensures that this tends to predominate. As it lacks one fatty acid in comparison to phosphatidic acid, it is a much more hydrophilic molecule, while the additional hydroxyl group strengthens hydrogen bonding within membranes, properties that may be important for its function in cells. Although lysophosphatidic acid is present at very low levels only in animal tissues, it is extremely important biologically, influencing many biochemical processes. It is a biosynthetic precursor of phosphatidic acid, but there is particular interest in its role as a lipid mediator with growth factor-like activities. For example, it is rapidly produced and released from activated platelets to influence target cells. Biosynthesis: In the circulation, the most important source of lysophosphatidic acid is the activity of an enzyme with lysophospholipase D-like activity and known as ‘autotaxin’ on lysophosphatidylcholine (200 μM in plasma) to yield LPA in an albumin-bound form mainly, although it is relatively soluble in aqueous media because of its polarity and small size. This lipid is more abundant in serum (1 to 5 μM) than in plasma (100 nM), because of the release of its main precursor, lysophosphatidylcholine, from activated platelets during coagulation. Autotaxin is a member of the nucleotide pyrophosphatase-phosphodiesterase family and is also present in cerebrospinal and seminal fluids and many other tissues including cancer cell lines from which it was first isolated and characterized. Indeed, the name derives from the finding that it promoted chemotaxis on melanoma cells in an autocrine fashion. It binds to target cells via integrin and heparan sulfate proteoglycans and this may assist the delivery of lysophosphatidic acid to its receptors. Genetic deletion of the enzyme in mice results in aberrant vascular and neuronal development and soon leads to death of the embryos. However, the overexpression of autotaxin causes physical defects also and is eventually lethal to embryos. Figure \(8\) shows the pathways for synthesis of lysophosphatidic acid. While autotaxin is the primary source of extracellular lysophosphatidic acid, it is now established that it is produced intracellularly by a wide variety of cell types by various mechanisms often with phosphatidic acid, derived from other phospholipids by the action of phospholipase D, as the primary precursor. For example, hydrolysis of phosphatidic acid by a phospholipase A2 (PLA2) is the main mechanism in platelets, but other cellular enzymes involved include a phosphatidic acid-selective phospholipase A1 (PLA1) producing sn-2-acyl-lysophosphatidic acid, a monoacylglycerol kinase (utilizing monoacylglycerols produced by the action of lipid phosphate phosphatases) and glycerol-3-phosphate acyltransferase (the first step in phosphatidic acid biosynthesis). In particular, secretory PLA2-IIA (sPLA2-IIA) is able to induce the release of LPA from phosphatidic acid exposed on the surface of extracellular vesicles derived from platelets and Ca2+-loaded erythrocytes upon stimulation by pro-inflammatory cytokines. General function: Although lysophospholipids are relatively small molecules, they carry a high content of information through the nature of the phosphate head group, the positional distribution of the fatty acids on the glycerol moiety, the presence of ether or ester linkages to the glycerol backbone, and the chain-length and degree and position of saturation of the fatty acyl chains. Lysophosphatidic acid acts upon nearly all cell types, often as a proliferative and pro-survival signal, inducing cellular invasion, migration and differentiation, while stimulating smooth muscle and fibroblast contraction, cytoskeletal rearrangement, secretion of cytokines/chemokines and numerous other effects. Many of these activities are displayed also by the 1-O-alkyl- and alkenyl-ether forms, which can be derived from platelet activating factor. On the other hand, it is possible that much of the lysophosphatidic acid produced intracellularly is used for synthesis of other phospholipids rather than for signaling purposes. Receptors: The informational content of the lysophosphatidic acid molecule leads to selectivity in the functional relationship with cell receptors. As most mammalian cells express receptors for lysophosphatidic acid, this lipid may initiate signaling in the cells in which it is produced, as well as affecting neighboring cells. Characterization of cloned lysophosphatidic acid receptors in combination with strategies of molecular genetics has allowed determination of both signaling and biological effects that are dependent on receptor mechanisms. At least six G protein-coupled receptors that are specific for lysophosphatidic acid have now been identified in vertebrates, each found in particular organs and coupled to at least one or more of the four heterotrimeric Gα proteins and designated LPAR1 to LPAR6, of which LPAR1 is virtually ubiquitous in tissues. These vary appreciably amino acid sequences but are classified into two subgroups, the EDG (LPAR1-3) and P2Y (LPAR4-6) families, with differing tissue distributions. Most cell types express these receptors in different combinations. There is also some interaction with transient receptor potential cation channel V1 (TRPV1), peroxisome proliferator-activated receptor gamma (PPARγ) and other proteins. Plasma lysophosphatidic acid binds to its receptors while it is bound to albumin. Experimental activation of the LPAR receptors has shown that a range of downstream signaling cascades are mediated by lysophosphatidic acid signaling via these various receptors. These include activation of adenylyl cyclase, cAMP production, intracellular Ca2+ and K+ production (by activating ion channels), protein kinases, phospholipase C, phosphatidylinositol 3-kinase, small GTPases (Ras, Rho, Rac), release of arachidonic acid, and much more. In this way, lysophosphatidic acid regulates cell survival, proliferation, cytoskeleton re-arrangement, motility, cytokine secretion, cell differentiation and many other vital cellular processes. Sometimes, lysophosphatidic acid appears to function in contradictory ways, and there is evidence that it is involved in cell survival in some circumstances and in programmed cell death in others, for example. Signaling by lysophosphatidic acid has regulatory functions in the mammalian reproductive system, both male and female, facilitating oocyte maturation and spermatogenesis through the action of the receptors LPAR1 to LPAR3. During early gestation, it regulates vascular remodeling at the maternal-fetal interface. There is also evidence that the lipid is involved in brain development, through its activity in neural progenitor cells, neurons, and glia, and in vascular remodeling. In the central nervous system, these receptors are thought to play a central role in both triggering and maintaining neuropathic pain by mechanisms that may involve demyelination of damaged nerves. Lysophosphatidic acid has been found in saliva in significant amounts, and it has been suggested that it is involved in wound healing in the upper digestive organs such as the mouth, pharynx, and oesophagus. When applied topically to skin wounds, it has similar effects probably by stimulating proliferation of new cells to seal the wound. Receptor LPAR6 together with the phospholipase A1 is required for the development of hair follicles, and this receptor is also involved in the regulation of endothelial blood-brain barrier function. The proliferation and survival of stem cells and their progenitors is regulated by lysophosphatidic acid signaling, while in bone cells, acting via LPAR1, lysophosphatidic acid is important for bone mineralization and repair. Disease: There is particular interest in the activity of lysophosphatidic acid in various disease states and cancer especially, as increased expression of autotaxin and the subsequent increased levels of lysophosphatidic acid have been reported in several primary tumors. For example, a finding that lysophosphatidic acid is markedly elevated in the plasma and peritoneal fluid (ascites) of ovarian cancer patients compared to healthy controls may be especially significant. Also, elevated plasma levels were found in patients in the first stage of ovarian cancer, suggesting that it may represent a useful marker for the early detection of the disease. It is believed that the secretory form of phospholipase A2 acts preferentially on lipids from damaged membranes or microvesicles, such as those produced by malignant cells, and this eventually results in increased levels of this lipid. Lysophosphatidic acid has been shown to stimulate the expression of genes for many different enzymes that lead to the proliferation of ovarian and other cancer cells and may induce cell migration via receptors LPAR1 to LPAR3 and possibly LPAR6, while LPAR4 and LPAR5 have opposing effects. Autotaxin and LPARs have been implicated in resistance to chemotherapy and radiation treatment in cancer therapy. As lysophosphatidic acid has growth-factor-like activities for many cell types that induce cell proliferation and migration, changes in cellular shape and increasing of endothelial permeability, it is perhaps not surprising that it is relevant to tumor biology. Treatment of various cancer cell types with lysophosphatidic acid promotes the expression and release of interleukin 8 (IL-8), which is a potent angiogenic factor, and thus it has a critical role in the growth and spread of cancers by enhancing the availability of nutrients and oxygen. There is evidence that signaling by lysophosphatidic acid is causally linked to hyperactive lipogenesis in cancer. For example, it activates the sterol regulatory element-binding protein (SREBP) together with the fatty acid synthase and AMP-activated protein kinase–ACC lipogenic cascades leading to elevated synthesis of lipids de novo. Increased autotaxin expression has been demonstrated in many different cancer cell lines, and the expression of many of the surface receptors for lysophosphatidic acid in cancer cells is aberrant. Cancer cells must evade the immune system during metastasis, and lysophosphatidic acid facilitates this process by inhibiting the activation of T cells. Therefore, lysophosphatidic acid metabolism is a target of the pharmaceutical industry in the search for new drugs for cancer therapy, aided by a knowledge of the crystal structures of three of the receptors. Signaling by lysophosphatidic acid has been implicated in many aspects of chronic inflammation, which it promotes by affecting the endothelium in several ways, for example by stimulating endothelial cell migration, the secretion of chemokines-cytokines and regulating the integrity of the endothelial barrier. Problems with lysophosphatidic acid signaling together with changes in autotaxin expression are believed to be factors in such metabolic and inflammatory disorders as obesity, insulin resistance, non-alcoholic fatty liver disease, rheumatoid arthritis, multiple sclerosis and cardiovascular disease. Further, there is evidence it contributes to neurological disorders, such as Alzheimer's disease and neuropathic pain, and to asthma, fibrosis and bone malfunction. Drugs that interact with the lysophosphatidic acid receptors are reported to be effective in attenuating symptoms of several diseases in animal models, and three have passed phase I and II clinical trials for idiopathic pulmonary fibrosis and systemic sclerosis in human patients. Drugs that target autotaxin production and catabolism of lysophosphatidic acid are also in development, and the steroidal anti-inflammatory agent, dexamethasone, appears to be especially useful. Under certain conditions, lysophosphatidic acid can become athero- and thrombogenic and might aggravate cardiovascular disease. As oxidized low-density lipoproteins promote the production of lysophosphatidic acid, its content in atherosclerotic plaques is high, suggesting that it might serve as a biomarker for cardiovascular disease. Indeed, lysophosphatidic acid promotes pro-inflammatory events that lead to the development of atheroma as well encouraging progression of the disease. By mediating platelet aggregation, it could lead to arterial thrombus formation. Related lipids: The sphingolipid analogue, sphingosine-1-phosphate, shows a similar range of activities to lysophosphatidic acid and the two lipids are often discussed together in the same contexts, although they may sometimes have opposing effects. Acute leukemia cells produce methyl-lysophosphatidic acids (the polar head-group is methylated). As these act as antigens to which a specific group of human T cells react strongly, it is possible that they might be a target for the immunotherapy of hematological malignancies. Other lysophospholipids are known to have distinctive biological functions. Catabolism: Deactivation of lysophosphatidic acid is accomplished by dephosphorylation to produce monoacylglycerols by a family of three lipid phosphate phosphatases (LPP1, 2 and 3), which also de-phosphorylate sphingosine-1-phosphate, phosphatidic acid and ceramide 1-phosphate in a non-specific manner. These are integral membrane proteins with the active site in the plasma membrane facing the extracellular environment, enabling them to access and hydrolyse extracellular lysophosphatidic acid and other phospholipids. Mice with a constitutive LPP3 deficiency are not viable, but this is not true for LPP1 and LPP2 knockout mice. Lysophosphatidic acid can be converted back to phosphatidic acid by a membrane-bound O-acyltransferase (MBOAT2) specific for lysophosphatidic acid (and lysophosphatidylethanolamine) with a preference for oleoyl-CoA as substrate. Phosphatidylcholine and Related Lipids Phosphatidylcholine - Structure and Occurrence Phosphatidylcholine or 1,2-diacyl-sn-glycero-3-phosphocholine (once given the trivial name 'lecithin') is a neutral or zwitterionic phospholipid over a pH range from strongly acid to strongly alkaline. It is usually the most abundant phospholipid in animals and plants, often amounting to almost 50% of the total complex lipids, and as such it is obviously a key building block of membrane bilayers. In particular, it makes up a very high proportion of lipids of the outer leaflet of the plasma membrane in animals. Virtually all the phosphatidylcholine in human erythrocyte membranes is present in the outer leaflet, for example, while in the plasma membranes of nucleated cells, 80 to 90% of this lipid is located on the outer leaflet. Phosphatidylcholine is also the principal phospholipid circulating in plasma, where it is an integral component of the lipoproteins, especially the HDL. On the other hand, it is less often found in bacterial membranes, perhaps ~10% of species, but there is none in the 'model' organisms Escherichia coli and Bacillus subtilis. In animal tissues, some of its membrane functions appear to be shared with the structurally related sphingolipid, sphingomyelin, although the latter has many unique properties of its own. Figure \(9\) shows the structure of phosphatidylcholine In animal tissues, phosphatidylcholine tends to exist in mainly in the diacyl form, but small proportions (in comparison to phosphatidylethanolamine and phosphatidylserine) of alkyl,acyl and alkenylacyl forms may also be present. As a generalization, animal phosphatidylcholine tends to contain lower proportions of arachidonic and docosahexaenoic acids and more of the C18 unsaturated fatty acids than the other zwitterionic phospholipid, phosphatidylethanolamine. Saturated fatty acids are most abundant in position sn-1, while polyunsaturated components are concentrated in position sn-2. Indeed, C20 and C22 polyenoic acids are exclusively in position sn-2, yet in brain and retina the unusual very-long-chain polyunsaturated fatty acids (C30 to C38) of the n-6 and n-3 families occur in position sn-1. Dietary factors obviously influence fatty acid compositions, but in comparing animal species, it would be expected that the structure of the phosphatidylcholine in the same metabolically active tissue would be somewhat similar in terms of the relative distributions of fatty acids between the two positions. Table \(1\) lists some representative data. Table \(1\). Positional distribution of fatty acids in the phosphatidylcholine of some animal tissues. Position Fatty acid 16:0 16:1 18:0 18:1 18:2 20:4 22:6 Rat liver [1] sn-1 23 1 65 7 1 trace sn-2 6 1 4 13 23 39 7 Rat heart [2] sn-1 30 2 47 9 11 - - sn-2 10 1 3 17 20 33 9 Rat lung [3] sn-1 72 4 15 7 3 - - sn-2 54 7 2 12 11 10 1 Human plasma [4] sn-1 59 2 24 7 4 trace - sn-2 3 1 1 26 32 18 5 Human erythrocytes [4] sn-1 66 1 22 7 2 - - sn-2 5 1 1 35 30 16 4 Bovine brain (gray matter) [5] sn-1 38 5 32 21 1 - - sn-2 33 4 trace 48 1 9 4 Chicken egg [6] sn-1 61 1 27 9 1 - - sn-2 2 1 trace 52 33 7 4 1, Wood, R. and Harlow, R.D. Arch. Biochem. Biophys., 131, 495-501 (1969); DOI. 2, Kuksis, A. et al. J. Lipid Res., 10, 25-32 (1969); DOI. 3, Kuksis, A. et al. Can. J. Physiol. Pharm., 46, 511-524 (1968); DOI. 4, Marai, L. and Kuksis, A. J. Lipid Res., 10, 141-152 (1969); DOI. 5, Yabuuchi, H. and O'Brien, J.S. J. Lipid Res., 9, 65-67 (1968); DOI. 6, Kuksis, A. and Marai, L. Lipids, 2, 217-224 (1967); DOI. There are some exceptions to the rule as the phosphatidylcholine in some tissues or organelles contains relatively high proportions of disaturated molecular species. For example, it is well known that lung phosphatidylcholine in most if not all animal species studied to date contains a high proportion (50% or more) of dipalmitoylphosphatidylcholine. The positional distributions of fatty acids in phosphatidylcholine in representative plants and yeast are listed in Table \(2\). In the leaves of the model plant Arabidopsis thaliana, saturated fatty acids are concentrated in position sn-1, but monoenoic fatty acids are distributed approximately equally between the two positions, and there is a preponderance of di- and triunsaturated fatty acids in position sn-2; the same is true for soybean ‘lecithin’. In the yeast Lipomyces lipoferus, the pattern is somewhat similar except that much of the 16:1 is in position sn-1. Table \(2\): Composition of fatty acids (mol %) in positions sn-1 and sn-2 in the phosphatidylcholine from plants and yeast. Position Fatty acid 16:0 16:1 18:0 18:1 18:2 18:3 Arabidopsis thaliana (leaves) [1] sn-1 42   4 5 23 26 sn-2 1   trace 5 47 47 Soybean 'lecithin' [2] sn-1 24   9 14 47 4 sn-2 5   1 13 75 6 Lipomyces lipoferus [3] sn-1 24 18 trace 37 16 4 sn-2 4 5 trace 39 31 19 1, Browse, J., Warwick, N., Somerville, C.R. and Slack, C.R. Biochem. J., 235, 25-31 (1986); DOI. 2, Blank, M.L., Nutter, L.J. and Privett, O.S. Lipids, 1, 132-135 (1966); DOI. 3, Haley, J.E. and Jack, R.C. Lipids, 9, 679-681 (1974); DOI. Phosphatidylcholine – Biosynthesis There are several mechanisms for the biosynthesis of phosphatidylcholine in animals, plants and micro-organisms. Choline itself is not synthesized as such by animal cells and is an essential nutrient, not only for phospholipid synthesis but also for cholinergic neurotransmission (acetylcholine synthesis) and as a source for methyl groups for numerous other metabolites. It must be obtained from dietary sources or by degradation of existing choline-containing lipids, for example those produced by the second pathway described below. Once taken across membranes and into cells by specific transporters, choline is immediately phosphorylated by a choline kinase (1) in the cytoplasm of the cell to produce phosphocholine, which is reacted with cytidine triphosphate (CTP) by the enzyme CTP:phosphocholine cytidylyltransferase (CCT) (2) to form cytidine diphosphocholine (CDP-choline). CTP + PC → CDP-choline + Pi The latter enzyme exists in two isoforms of which CCTα is the more important and is a soluble protein found first in the nucleoplasm, but then in the nucleoplasmic reticulum. This is considered to be the rate-limiting step in phosphatidylcholine biosynthesis, and the activity of the enzyme is regulated by signals from a sensor in the membrane that reports on the relative abundance of the final product. However, choline kinase (ChoKα) also has regulatory functions. Figure \(10\) shows an interactive iCn3D model of the mammalian (rat) CTP: Phosphocholine cytidylyltransferase catalytic domain (3HL4). Figure \(10\): Mammalian (rat) CTP-Phosphocholine cytidylyltransferase catalytic domain (3HL4). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...tBYRYk7vPKD7f6 The biologically active homodimer is shown. The B chain is colored by secondary structure and the A chain is shown in gray. Key active site residues (only shown in the A chain) are in CPK-colored sticks and labeled. A bound CDP-choline analog ((2-cytidylate-O'-phosphonyloxy)-ethyl-trimethylammonium) is shown in spacefill CPK colors. This enzyme (CCT) catalyzes the key regulatory and rate-limiting step in PC synthesis. The C-terminal domain binds membrane lipids and regulates the enzyme. The iCn3D model above is for the catalytic domain with the regulatory domain deleted. Two nonconserved active site side chains, His 168 and Tyr 173 interact with and position the phosphocholine. Other active site residues include Arg-196 in L6, Lys-122 in L2, and Asp-94. Figure \(11\) shows a simplified mechanism for the reaction In plants, nematodes and certain parasites, most phosphocholine is synthesized by sequential methylation of phosphoethanolamine by phospho-base N‑methyltransferases, but phosphatidylethanolamine is only methylated in this way in a few plant species. This is also the main route to free choline and betaine in plants. The CDP-choline produced is acted upon by the membrane-bound enzyme CDP-choline:1,2-diacylglycerol choline/ethanolamine-phosphotransferase in the endoplasmic reticulum (CEPT1), and a related choline phosphotransferase 1 (CPT1) in the trans-Golgi, which catalyse the reaction with sn-1,2-diacylglycerols to form phosphatidylcholine. The first of these is responsible for most phosphatidylcholine biosynthesis but with a somewhat different molecular species composition from the second, which has a preference for 1-alkyl precursors. This is the main pathway for the synthesis of phosphatidylcholine in animals and plants, and it is analogous to that for a major route to phosphatidylethanolamine; it is also found in a few bacterial species (e.g. Sinorhizobium meliloti). Phosphatidylcholine in mitochondria is obtained by transfer from the endoplasmic reticulum. Figure \(12\) shows the main pathways for PC synthesis in plants and animals. The discovery of the importance of this pathway depended a little on serendipity in that in experiments in the laboratory of Professor Eugene Kennedy, samples of adenosine triphosphate (ATP) contained some cytidine triphosphate (CTP) as an impurity. However, luck is of little value without receptive minds, and Kennedy and co-workers demonstrated that the impurity was an important metabolite that was essential for the formation of phosphatidylcholine. The above reaction, together with the biosynthetic mechanism for phosphatidylethanolamine, is significantly different from that for phosphatidylglycerol, phosphatidylinositol and cardiolipin. Both make use of nucleotides, but with the latter, the nucleotide is covalently linked directly to the lipid intermediate, i.e., cytidine diphosphate diacylglycerol. However, a comparable pathway to the latter for biosynthesis of phosphatidylcholine occurs in bacteria (see below). The source of the sn-1,2-diacylglycerol precursor, which is also a key intermediate in the formation of phosphatidylethanolamine and phosphatidylserine, and of triacylglycerols, is phosphatidic acid. In this instance, the important enzyme is phosphatidic acid phosphatase (also known as lipin or phosphatidate phosphatase’ or ‘lipid phosphate phosphatase’ or ‘phosphatidate phosphohydrolase’). Figure \(13\) shows the biosynthesis of the diacyl precursor of PC This enzyme is also important for the production of diacylglycerols as essential intermediates in the biosynthesis of triacylglycerols and of phosphatidylethanolamine. Yeasts contain two such enzymes, one of which is Mg2+-dependent (PAP1) and the other Mg2+-independent (PAP2). In mammals, much of the phosphatidic acid phosphatase activity resides in three related cytoplasmic proteins, termed lipins-1, -2, and -3. Lipin-1 is found mainly in adipose tissue, while lipin-2 is present mainly in liver. They are unique among biosynthetic enzymes for glycerolipids in that they can transit among cellular membranes rather than remain tethered to membranes. Of these lipin-1 is most important and exists in three isoforms, lipin-1α, lipin-1β and lipin-1γ with lipin-1α located mainly in the nucleus and lipin-1β in the cytoplasm. Lipin-1γ is present primarily in brain. The second pathway for biosynthesis of phosphatidylcholine involves sequential methylation of phosphatidylethanolamine, with S-adenosylmethionine (SAM) as the source of methyl groups, with mono- and dimethylphosphatidylethanolamine as intermediates and catalyzed by the enzyme phosphatidylethanolamine N‑methyltransferase. A single enzyme (~20 KDa) in two isoforms catalyses all three reactions in hepatocytes; the main form is located in the endoplasmic reticulum (ER) where it spans the membrane, while the second is found in the mitochondria-associated ER membrane. At least two N-methyltransferases are present in yeasts. This is a major pathway in the liver, generating one third of the phosphatidylcholine in this organ, but not in other animal tissues or in general in higher organisms. It may be the main route to phosphatidylcholine in those bacterial species that produce this lipid and in yeasts, but it appears to operate in only a few species of higher plants. When choline is deficient in the diet, this liver pathway is especially important. Figure \(14\) shows the synthesis of PC via methylation of PE. A by-product of the biosynthesis of phosphatidylcholine from phosphatidylethanolamine is the conversion of S‑adenosylmethionine to S‑adenosylhomocysteine, which is hydrolyzed in the liver to adenosine and homocysteine. An elevated level of the latter in plasma is a risk factor for cardiovascular disease and myocardial infarction. Phosphatidylcholine biosynthesis by both pathways in the liver is necessary for normal secretion of the plasma lipoproteins (VLDL and HDL), and it is relevant to a number of human physiological conditions. It should be noted that all of these pathways for the biosynthesis of diacylphosphatidylcholine are very different and are separated spatially from that producing alkyl, acyl- and alkenylacyl-phosphatidylcholines de novo. Also, synthesis of phosphatidylcholine does not occur uniformly throughout the endoplasmic reticulum but is located at membrane interfaces or where it meets other organelles, and especially where the membrane is expanding dynamically. The enzymes in the endoplasmic reticulum responsible for the synthesis of all phospholipids are orientated in such a manner that their active sites are exclusively facing the cytosol. Problems would arise if there were a rapid expansion of the cytosolic leaflet while the luminal leaflet did not change, but a phospholipid transporter known as a scramblase enables a rapid bidirectional flip-flop of phospholipids between leaflets of the bilayer in an energy-independent manner. Compositional asymmetry in first seen in the trans-Golgi and is completed before the plasma membrane is formed with phosphatidylcholine and sphingolipids present mainly in the exofacial (outer) leaflet while phosphatidylethanolamine and phosphatidylserine are enriched in the cytosolic leaflet. Dietary phosphatidylcholine is rapidly hydrolyzed in the proximal small intestine by pancreatic enzymes with formation of lysophosphatidylcholine (and free fatty acids). Further hydrolysis can occur in the jejuno-ileal brush-border by the action of the membrane phospholipases, with the release of glycerophosphocholine, but much of the lysophosphatidylcholine is reacylated by the lyso-PC-acyl-CoA-acyltransferase 3 for export in chylomicrons. In plant cells, phosphatidylcholine biosynthesis occurs mainly in the endoplasmic reticulum, and it is a major component of most membranes other than the internal membranes of plastids; it is absent from the thylakoids and the inner envelope membrane, but is the main glycerolipid of the outer monolayer of the outer envelope membrane. Further complications arise in plants in that turnover or partial synthesis via lysophosphatidylcholine occurs in different organelles from different fatty acid pools or with enzymes with differing specificities, and in addition, fatty acids esterified to phosphatidylcholine serve as substrates for desaturases. The result is that an appreciable pool of the diacylglycerols for the biosynthesis of triacylglycerols, galactosyldiacylglycerols, and other glycerolipids pass through phosphatidylcholine as an intermediate so that the fatty acid compositions in different membranes change after the initial synthetic process. This mechanism has obvious differences from the remodeling of molecular species in animal tissues discussed next, although a comparable exchange of acyl groups does occur in part catalyzed by acyl transferases (see next section). Some transfer of phosphatidylcholine per se from the endoplasmic reticulum to plastids may occur via contact points between the two membranes or may be facilitated by specific transport proteins. While phosphatidylcholine is a major lipid in yeasts, recent work suggests that it is not essential if suitable alternative growth substrates are available, unlike higher organisms where perturbation of phosphatidylcholine synthesis can lead to inhibition of growth or even cell death. Remodeling of Phosphatidylcholine - the Lands' cycle Whatever the mechanism of biosynthesis of phosphatidylcholine in animal tissues, it is apparent that the fatty acid compositions and positional distributions on the glycerol moiety are determined post synthesis by extensive re-modeling involving orchestrated reactions of hydrolysis (phospholipase A2 mainly) to lysophosphatidylcholine, acyl-CoA synthesis and re-acylation by lysophospholipid acyltransferases or transacylases, a series of reactions that is sometimes termed the 'Lands' Cycle' after its discoverer W.E.M. (Bill) Lands. Similar processes occur with all glycerophospholipid classes. The final composition of the lipid is achieved by a mixture of synthesis de novo and the remodeling pathway. There are at least fifteen different groups of enzymes in the phospholipase A2 super-family, which differ in calcium dependence, cellular location and structure. All hydrolyze the sn-2 ester bond of phospholipids specifically, generating a fatty acid and lysophospholipid, both of which have important functions in their own right in addition to their role in the Lands cycle. There is also a phospholipase A1 family of enzymes, which are esterases that are able to cleave the sn-1 ester bond but are less important in this context. Figure \(15\) shows Land's cycle The re-acylation step is catalyzed by membrane-bound coenzyme A-dependent lysophosphatidylcholine acyltransferases such as LPCAT3 (also designated ‘MBOAT5’), which is located chiefly within the endoplasmic reticulum, though also in mitochondria and the plasma membrane in organs such as the liver, adipose tissue and pancreas. It maintains systemic lipid homeostasis by regulating lipid absorption and composition in the intestines, the secretion of lipoproteins, and lipogenesis de novo in liver, and is notable in that it incorporates linoleoyl and arachidonoyl chains specifically into lysophosphatidylcholine (as does a related enzyme LPCAT2). There is also a CoA-independent acyltransferase in inflammatory cells that transfers arachidonic acid from phosphatidylcholine to ethanolamine-containing phospholipids. While LPCAT3 prefers 1-acyl lysophosphatidylcholine as an acyl acceptor, LPCAT2 utilizes both 1-acyl and 1-alkyl precursors. LPCAT2 is highly expressed in inflammatory cells such as macrophages and neutrophils, which contain ether-phospholipids, where it contributes to the production of eicosanoid lipid mediators. The highly saturated molecular species of phosphatidylcholine found in lung surfactant are formed from species with a more conventional composition by remodeling by an acyltransferase with a high specificity for palmitoyl-CoA acid (LPCAT1). In other tissues, those species containing high proportions of polyunsaturated fatty acids depend more on synthesis de novo. These and further related enzymes are involved in remodeling of all other phospholipids. Over-expression of the genes for these enzymes is associated with the progression of many different cancers and may be involved in other pathological conditions. Phosphatidylcholine has a central role in glycerolipid metabolism in plants and remodeling occurs for reasons and by mechanisms that are rather different from those in animal cells as described briefly above. For example, there is extensive remodeling as a site of fatty acid desaturation and as the main entry point for acyl groups exported from the plastid into the endoplasmic reticulum. In addition, the remodeling of phosphatidylcholine provides fatty acids for triacylglycerol synthesis in developing seeds and diacylglycerols for the synthesis of thylakoid lipids such as galactosyldiacylglycerols. In Arabidopsis, two lysophosphatidylcholine acyltransferases, LPCAT1 and LPCAT2, are involved in remodeling in developing seeds and leaves, with some preference for position sn-2 using fatty acids exported from the plastid. In some plant species, there is a strong preference for C18‑unsaturated acyl chains over 16:0. However, the lipases that generate lysophosphatidylcholine from phosphatidylcholine for this purpose are not yet known. Some remodeling in plant membranes occurs in response to stress. The yeast Saccharomyces cerevisiae is able to reacylate glycerophosphocholine, generated endogenously by the action of phospholipase B (an enzyme with both phospholipase A1 and A2 activities) on phosphatidylcholine, with acyl-CoA in the microsomal membranes by means of a glycerophosphocholine acyltransferase (Gpc1) to produces lysophosphatidylcholine, which can be converted back to phosphatidylcholine by the lysophospholipid acyltransferase (Ale1) with appreciable changes in the molecular species composition. The process is regulated in coordination with the other main lipid pathways and affects yeast growth. The enzyme Gpc1 does not affect other phospholipids in yeasts. A similar mechanism appears to operate in some plant species. Figure \(16\) shows variants of the Land's cycle. Catabolism Phosphatidylcholine (and most other glycerophospholipids) in membranes can be metabolized by lipolytic enzymes, especially phospholipases, some isoforms of which are specific for particular lipid classes in humans. For example, in addition to the action of phospholipase A (discussed above), phospholipase C (six families in mammals differing in expression and subcellular distribution) yields diacylglycerols together with phosphocholine by hydrolyzing glycerophospholipids at the phosphodiester bond, a process that is especially important in relation to phosphoinositide metabolism. The sphingomyelin synthases also have phospholipase C activity (in the absence of ceramide). Phospholipase D generates phosphatidic acid and choline, while phospholipase B removes both fatty acids to yield glycerophosphocholine. Figure \(17\) shows the activities of phospholipases on phosphatidyl choline. On catabolism in this way, the lipid components are re-cycled or they may have signaling functions, while much of the choline is re-used for phosphatidylcholine biosynthesis, often after being returned to the liver (the CDP-choline cycle). Some choline is oxidized in the kidney and liver to betaine, which serves as a donor of methyl groups for S-adenosylmethionine production, and some is lost through excretion of phosphatidylcholine in bile. A proportion is used in nervous tissues for production of acetylcholine, a neurotransmitter of importance to learning, memory and sleep. Phosphatidylcholine in the high-density lipoproteins of plasma is taken up by the liver, and perhaps surprisingly a high proportion of this is eventually converted to triacylglycerols via diacylglycerol intermediates. Phosphatidylcholine – Biological Functions Because of the generally cylindrical shape of the molecule, phosphatidylcholine organizes spontaneously into bilayers, so it is ideally suited to serve as the bulk structural element of biological membranes, and as outlined above it is makes up a high proportion of the lipids in the outer leaflet of the plasma membrane. The unsaturated acyl chains are kinked and confer fluidity on the membrane. Such properties are essential to act as a balance to those lipids that do not form bilayers or that form specific micro-domains such as rafts. While phosphatidylcholine does not induce curvature of membranes, as may be required for membrane transport and fusion processes, it can be metabolized to form lipids that do. In contrast, dipalmitoyl phosphatidylcholine is the main surface-active component of human lung surfactant, although in other animals the lung surfactant can be enriched in some combination of short-chain disaturated and monounsaturated species, mainly palmitoylmyristoyl- and palmitoylpalmitoleoyl- in addition to the dipalmitoyl-lipid. This is believed to provide alveolar stability by decreasing the surface tension at the alveolar surface to a very low level during inspiration while preventing alveolar collapse at the end of expiration. Also, the internal lipids of the animal cell nucleus (after the external membrane has been removed) contain a high proportion of disaturated phosphatidylcholine. This is synthesized entirely within the nucleus, unlike phosphatidylinositol for example, and in contrast to other cellular lipids its composition cannot be changed by extreme dietary manipulation; it has been suggested that it may have a role in stabilizing or regulating the structure of the chromatin, as well as being a source of diacylglycerols with a signaling function. A further unique molecular species, 1-oleoyl-2-palmitoyl-phosphatidylcholine, is located specifically at the protrusion tips of neuronal cells and appears to be essential for their function, while 1-palmitoyl-2-arachidonoyl-phosphatidylcholine is important in the regulation of the progression of the cell cycle and cell proliferation, and this is independent of eicosanoid production. Phosphatidylcholine is present bound non-covalently in the crystal structures of a number of membrane proteins, including cytochrome c oxidase and yeast cytochrome bc1. The ADP/ATP carrier protein has two binding sites for phosphatidylcholine, one on each side. In addition, it is known that the enzyme 3‑hydroxybutyrate dehydrogenase must be bound to phosphatidylcholine before it can function optimally. Both the head group and the acyl chains may be involved in the interactions depending on the protein. As noted above, phosphatidylcholine is by far the most abundant phospholipid component in plasma and in all plasma lipoprotein classes. Although it is especially abundant in high density lipoproteins (HDL), it influences strongly the levels of all circulating lipoproteins and especially of the very-low-density lipoproteins (VLDL), which are surrounded by a phospholipid monolayer. Indeed, phosphatidylcholine with polyunsaturated fatty acids in position sn-2 is essential for the assembly and secretion of VLDLs and chylomicrons in liver and the intestines, and it must be synthesized de novo in the latter. Similarly, phosphatidylcholine synthesis is required to stabilize the surface of lipid droplets in tissues where triacylglycerols are stored. Some of the phosphatidylcholine synthesized in the liver is secreted into bile by a specific flippase together with bile acids where it assists in the emulsification of dietary triacylglycerols in the intestinal lumen to facilitate their hydrolysis and uptake. Eventually, it is absorbed across the intestinal brush border membrane after hydrolysis to lysophosphatidylcholine, which may then be involved in the initiation of chylomicron formation in the endoplasmic reticulum of enterocytes by activation of a protein kinase. In addition, phosphatidylcholine produced in enterocytes is secreted into the intestinal lumen and forms part of the hydrophobic mucus layer that protects the intestinal surface. Phosphatidic acid generated from phosphatidylcholine by the action of phospholipase D in plants has key signaling functions. Similarly, phosphatidic acid generated in this way from phosphatidylcholine in animals is involved in the metabolism and signaling function of phosphoinositides by activating phosphatidylinositol 4-phosphate 5-kinase, the main enzyme generating the lipid second messenger phosphatidylinositol-4,5-bisphosphate. The plasmalogen form of phosphatidylcholine may also have a signaling function, as thrombin treatment of endothelial cells activates a selective hydrolysis (phospholipase A2) of molecular species containing arachidonic acid in the sn-2 position, releasing this fatty acid for eicosanoid production, while the diacyl form of phosphatidylcholine may have a related function in signal transduction in other tissues. In addition, phosphatidylcholine may have a role in signaling via the generation of diacylglycerols by phospholipase C, especially in the nucleus. Although the pool of the precursor is so great in many tissues that turnover is not easily measured, the presence of phospholipases C and D that are specific for phosphatidylcholine and are activated by a number of agonists suggests such a function especially in the cell nucleus. Diacylglycerols formed in this way would be much more saturated than those derived from phosphatidylinositol, and would not be expected to be as active in some functions. Phosphatidylcholine is the biosynthetic precursor of sphingomyelin and as such must have some influence on the many metabolic pathways that constitute the sphingomyelin cycle. It is also a precursor for phosphatidic acid, lysophosphatidylcholine and platelet-activating factor, each with important signaling functions, and of phosphatidylserine. Because of the increased demand for membrane constituents, there is enhanced synthesis of phosphatidylcholine in cancer cells and solid tumours; the various biosynthetic and catabolic enzymes are seen as potential targets for the development of new therapeutic agents. Impaired phosphatidylcholine biosynthesis has been observed in a number of pathological conditions in the liver in humans, including the development of non-alcoholic fatty liver disease, liver failure and impaired liver regeneration. Similarly, a deficiency in phosphatidylcholine or an imbalance in the ratio of phosphatidylcholine to phosphatidylethanolamine has negative effects upon insulin sensitivity and glucose homeostasis in skeletal muscle. Plants and bacteria: In addition to its structural role in plant membranes, phosphatidylcholine levels at the shoot apex correlate with flowering time, and this lipid is believed to bind to the Flowering Locus T, a master regulator of flowering. Molecular species containing relatively low levels of α-linolenic acid are involved. Diacylglycerols formed by the action of a family of enzymes of the phospholipase C type on phosphatidylcholine, as opposed to phosphatidylinositol, may be more important in plants and especially during phosphate deprivation for the generation of precursors for galactolipid biosynthesis and perhaps for lipid re-modeling more generally. In prokaryotes, phosphatidylcholine is essential for certain symbiotic and pathogenic microbe-host interactions. For example, in human pathogens such as Brucella abortus and Legionella pneumophila, this lipid is necessary for full virulence, and the same is true for plant pathogens, such as Agrobacterium tumefaciens. Bacteria symbiotic with plants, e.g. the rhizobial bacterium Bradyrhizobium japonicum, require it to establish efficient symbiosis and root nodule formation. Lysophosphatidylcholine Figure \(18\) shows the structure of lysophosphatidylcholine Lysophosphatidylcholine (LPC), with one mole of fatty acid per mole of lipid in position sn-1, is found in trace amounts in most animal tissues, although there are relatively high concentrations in plasma (150–500µM). It is produced by hydrolysis of dietary and biliary phosphatidylcholine and is absorbed as such in the intestines, but it is re-esterified before being exported in the lymph. In addition, it is formed in most tissues by hydrolysis of phosphatidylcholine by means of the superfamily of phospholipase A2 enzymes as part of the de-acylation/re-acylation cycle that controls the overall molecular species composition of the latter, as discussed above. Much of the LPC in the plasma of animal species is secreted by hepatocytes into plasma in a complex with albumin, but an appreciable amount is formed in plasma by the action of the enzyme lecithin:cholesterol acyltransferase (LCAT), which is secreted from the liver. This catalyses the transfer of fatty acids from position sn-2 of phosphatidylcholine to free cholesterol in plasma, with formation of cholesterol esters and of course of lysophosphatidylcholine, which consists of a mixture of molecular species with predominately saturated and mono- and dienoic fatty acid constituents. Some LPC is formed by the action of an endothelial lipase on phosphatidylcholine in HDL. At high concentrations, lysophosphatidylcholine can disrupt membranes, while some biological effects at low concentrations may be simply due to its ability to diffuse readily into membranes, altering their curvature and indirectly affecting the properties of membrane proteins. In plasma, it is bound to albumin and lipoproteins so that its effective concentration is reduced to a relatively safe level. Lysophosphatidylcholine is considered to be an important factor in cardiovascular and neurodegenerative diseases. It is usually considered to have pro-inflammatory properties and it is known to be a pathological component of oxidized lipoproteins (LDL) in plasma and of atherosclerotic lesions, when it is generated by over-expression or enhanced activity of phospholipase A2. Its concentration is elevated in joint fluids from patients with rheumatoid arthritis. In addition, it is a major component of platelet-derived microvesicles and activates a specific receptor in platelets that ultimately leads to vascular inflammation, increasing the instability of atherosclerotic plaques. The intracellular acyltransferase LPCAT cannot remove lysophosphatidylcholine directly from plasma or lipoproteins, nor do there appear to be any enzymes with lysophospholipase A1 activity in the circulation. Lysophosphatidylcholine blocks the formation of early hemifusion intermediates required for cell-cell fusions. Lysophosphatidylcholine in insect bites attracts inflammatory cells to the site, enhances parasite invasion, and inhibits the production of nitric oxide, for example in Chagas disease. Elevated levels of 26:0‑lysophosphatidylcholine in blood are reported to be characteristic of Zellweger spectrum disorders (the result of a defect in peroxisome biogenesis). Elevated levels of lysophosphatidylcholine have been identified in cervical cancer and may be diagnostic for the disease. On the other hand, reduced concentrations of lysophosphatidylcholine are observed in some malignant cancers, and it has protective effects in patients undergoing chemotherapy. Stearoyl-lysophosphatidylcholine has an anti-inflammatory role in that it is protective against lethal sepsis in experimental animals by various mechanisms, including stimulation of neutrophils to eliminate invading pathogens through a peroxide-dependent reaction. Similarly, there are reports that lysophosphatidylcholine may have beneficial effects in rheumatoid arthritis and a number of other diseases. However, there are suggestions that some experimental studies in vitro of the activity of lysophosphatidylcholines may be flawed because insufficient levels of carrier proteins were used. A further point for consideration is that lysophosphatidylcholine is the precursor of the key lipid mediator lysophosphatidic acid via the action of the enzyme autotaxin in plasma, and this may be the true source of some of the effects described for the former, especially on cell migration and survival. There is evidence to suggest that lysophosphatidylcholine containing docosahexaenoic (DHA) and eicosapentaenoic (EPA) acids, presumably in position sn-2, in plasma targets more of these fatty acids into the brain, via a specific receptor/transporter at the blood-brain barrier known as the sodium-dependent LPC symporter 1 (MFSD2A), than occurs from the corresponding fatty acids in unesterified form. Hepatic lipase is especially important for generation of these lipids. This finding is now being explored in relation to potential therapeutic applications for neurological diseases, cognitive decline and dementia. Similarly, at the maternal plasma/placental interface, phosphatidylcholine is taken up and hydrolyzed to sn‑2‑lysophosphatidylcholine, presumably by the endothelial lipase, to facilitate transfer of polyunsaturated fatty acids across the basal membrane into the fetal circulation with the aid of the same LPC transporter. Lysophosphatidylcholine has been found to have some functions in cell signaling , and specific receptors (coupled to G proteins) have been identified, i.e., GPR119, GPR40 and GPR55. It activates the specific phospholipase C that releases diacylglycerols and inositol triphosphate with resultant increases in intracellular Ca2+ and activation of protein kinase C. Increased glucose-stimulated insulin secretion has been observed in different cell systems. Lysophosphatidylcholine also activates the mitogen-activated protein kinase in certain cell types, and it promotes demyelination in the nervous system. By interacting with the TRPV4 ion channels of skin keratinocytes, it causes persistent itching. Identification of a highly specific phospholipase A2γ in peroxisomes that is unique in generating sn-2-arachidonoyl lysophosphatidylcholine suggests that this may be of relevance to eicosanoid generation and signaling . For example, there is reportedly an enrichment of 2-arachidonoyl-lysophosphatidylcholine in carotid atheroma plaque from type 2 diabetic patients. In vascular endothelial cells, it induces the important pro-inflammatory mediator cyclooxygenase-2 (COX-2), a key enzyme in prostaglandin synthesis. However, it has beneficial effects on the innate immune system as it is able to activate macrophages and increase their phagocytic activity in the presence of T lymphocytes. As lysophospholipids in general and lysophosphatidylcholine in particular are important signaling molecules within mammalian cells, their levels are closely regulated, mainly by the action of the lysophospholipases A1 and A2 (LYPLA1 and LYPLA2), depending on the position to which the fatty acid is esterified; these are cytosolic serine hydrolases with esterase and thioesterase activity. The glycerophosphocholine produced can enter the Lands' cycle or be further degraded. In relation to plants, amylose-rich starch granules of cereal grains contain lysophosphatidylcholine as virtually the only lipid in the form of inclusion complexes or lining channels in the starch macromolecules. Phosphatidylethanolamine and Related Lipids Phosphatidylethanolamine – Structure and Occurrence Phosphatidylethanolamine or 1,2-diacyl-sn-glycero-3-phosphoethanolamine (once given the trivial name 'cephalin') is usually the second most abundant phospholipid in animal and plant lipids, after phosphatidylcholine, and it is frequently the main lipid component of microbial membranes. It can amount to 20% of liver phospholipids and as much as 45% of those of brain; higher proportions are found in mitochondria than in other organelles. As such, it is obviously a key building block of membrane bilayers, and it is present exclusively in the inner leaflet of the plasma membrane in animal cells, for example. It is a neutral or zwitterionic phospholipid (at least in the pH range 2 to 7), with the structure shown (with one specific molecular species illustrated as an example). Figure \(19\) shows the structure of phosphatidylethanolamine. In animal tissues, phosphatidylethanolamine tends to exist in diacyl, alkyl,acyl and alkenyl, acyl forms. As much as 70% of the phosphatidylethanolamine in some cell types (especially inflammatory cells, neurons and tumor cells) can have an ether linkage, but in liver, the plasmalogen form of phosphatidylethanolamine, i.e., with an O‑alk-1’-enyl linkage, accounts for only 0.8% of total phospholipids. Generally, there is a much higher proportion of phosphatidylethanolamine with ether linkages than of phosphatidylcholine. If biosynthesis of the plasmalogen form is inhibited by physiological conditions, it is replaced by the diacyl form so that the overall content of the phospholipid remains constant. In general, animal phosphatidylethanolamine tends to contain higher proportions of arachidonic and docosahexaenoic acids than the other zwitterionic phospholipid, phosphatidylcholine. These polyunsaturated components are concentrated in position sn-2 with saturated fatty acids most abundant in position sn-1, as illustrated for rat liver and chicken egg in Table \(3\). In most other species, it would be expected that the structure of the phosphatidylethanolamine in the same metabolically active tissues would exhibit similar features. Table \(3\): Positional distribution of fatty acids in phosphatidylethanolamine in animal tissues. Position Fatty acid 14:0 16:0 18:0 18:1 18:2 20:4 22:6 Rat liver [1] sn-1   25 65 8 sn-2 2 11 8 8 10 46 13 Chicken egg [2] sn-1   32 59 7 1 sn-2   1 1 25 22 29 12 1, Wood, R. and Harlow, R.D., Arch. Biochem. Biophys., 131, 495-501 (1969); 2, Holub, B.J. and Kuksis, A. Lipids, 4, 466-472 (1969); The O-alkyl and O-alkenyl chains at the sn-1 position of the analogous ether lipids generally consist of 16:0, 18:0 or 18:1 chains, whereas arachidonic and docosahexaenoic acids are the most abundant components at the sn-2 position. The positional distributions of fatty acids in phosphatidylethanolamine from the leaves of the model plant Arabidopsis thaliana are listed in Table \(4\). Here also saturated fatty acids are concentrated in position sn-1, and there is a preponderance of di- and triunsaturated in position sn-2. The pattern is somewhat different for the yeast Lipomyces lipoferus, where the compositions of the two positions are relatively similar. Table \(4\)​​​​​​​: Composition of fatty acids (mol %) in positions sn-1 and sn-2 in the phosphatidylethanolamine from leaves of Arabidopsis thaliana and from Lipoferus lipoferus . Position Fatty acid 16:0 16:1 18:0 18:1 18:2 18:3 A. thaliana [1] sn-1 58 trace 4 5 15 18 sn-2 trace trace trace 2 60 38 L. lipoferus [2] sn-1 29 18 4 28 13 6 sn-2 23 15 3 34 17 6 1, Browse, J., Warwick, N., Somerville, C.R. and Slack, C.R. Biochem. J., 235, 25-31 (1986); DOI. 2, Haley, J.E. and Jack, R.C. Lipids, 9, 679-681 (1974); DOI . Phosphatidylethanolamine – Biosynthesis The two main pathways employed by mammalian cells for the biosynthesis of phosphatidylethanolamine are the CDP-ethanolamine pathway, i.e., one of the general routes to phospholipid biosynthesis de novo in plants and animals, and the phosphatidylserine decarboxylase pathway, which occur in two spatially separated organelles - the endoplasmic reticulum and mitochondria, respectively. Ethanolamine is obtained by decarboxylation of serine in plants, and in animals most must come from dietary sources and requires facilitated transport into cells. A small amount of ethanolamine phosphate comes from catabolism of sphingosine-1-phosphate, and this is essential for the survival of the protozoon Trypanosoma brucei. The initial steps in phosphatidylethanolamine biosynthesis occur in the cytosol with first the phosphorylation of ethanolamine by two specific ethanolamine kinases to produce ethanolamine phosphate; the reverse reaction can occur by means of the enzyme ethanolamine phosphate phosphorylase and this may have a regulatory function in some tissues. The second step is rate-limiting, i.e., reaction of the product with cytidine triphosphate (CTP) to form cytidine diphosphoethanolamine catalyzed by CTP:phosphoethanolamine cytidylyltransferase. In the final step, a membrane-bound enzyme CDP-ethanolamine:diacylglycerol ethanolaminephosphotransferase catalyses the reaction of cytidine diphosphoethanolamine with diacylglycerol to form phosphatidylethanolamine. There are two such enzymes, ethanolamine phosphotransferase 1 (EPT1) in the Golgi and choline/EPT1 (CEPT1) in the endoplasmic reticulum, but EPT1 is more important for the biosynthesis of the plasmalogen form, 1-alkenyl-2-acyl-glycerophosphoethanolamine, and especially molecular species containing polyunsaturated fatty acids, while CEPT1 produced species with shorter-chain fatty acids. The diacylglycerol precursor is formed from phosphatidic acid via the action of the enzyme phosphatidic acid phosphohydrolase. Figure \(20\) shows the synthesis of phosphatidyethanolamine in the ER/Golgi. The second major pathway is the conversion (decarboxylation) of phosphatidylserine to form phosphatidylethanolamine in mitochondria. Conservation of the this pathway from bacteria to humans suggests that it has been preserved to optimize mitochondrial performance. Mitochondria are not connected with the rest of the cell's membrane network by classical vesicular routes, but must receive and export small molecules through the nonvesicular transport at zones of close proximity with other organelles at membrane contact sites, such as a specific domain of the endoplasmic reticulum termed the mitochondria-associated membrane (MAM). Lipid transport of phosphatidylserine is then enabled by tethers that bridge two membranes, lipid transfer proteins and recruitment proteins. In this process, the lipid must traverse two aqueous compartments, the cytosol and the mitochondrial intermembrane space, to reach the inner mitochondrial membrane. The phosphatidylserine decarboxylase is located on the external aspect of the mitochondrial inner membrane, and most of the phosphatidylethanolamine in mitochondria is derived from this pathway. While various isoforms of the phosphatidylserine decarboxylation exist in prokaryotes, yeasts and mammals, the main forms designated 'PSD1' are found only in mitochondria and are related structurally. An isoform designated 'PSD2' is located in the endosomal membranes of yeasts, and the phosphatidylethanolamine formed is the preferred substrate for phosphatidylcholine biosynthesis. It is evident that cellular concentrations of phosphatidylethanolamine and phosphatidylserine are closely related and tightly regulated. Figure \(21\) shows the synthesis of phosphatidyethanolamine in mitochondria. In prokaryotic cells, such as E. coli, in which phosphatidylethanolamine is the most abundant membrane phospholipid, all of it is derived from phosphatidylserine decarboxylation. In this instance, the enzyme undergoes auto-cleavage for activation and utilizes a pyruvoyl moiety to form a Schiff base intermediate with phosphatidylserine to facilitate decarboxylation. This pathway is also important in mammalian cells and yeasts, although the relative contributions of the two main pathways for phosphatidylethanolamine synthesis in mammalian cells appears to depend on the cell type. In cells in culture, more than 80% of the phosphatidylethanolamine is reported to be derived from the phosphatidylserine decarboxylase pathway, but in hamster heart and rat hepatocytes, only ~5% of phosphatidylethanolamine synthesis comes from this route and most is from the CDP-ethanolamine pathway. In yeasts, 70% of the phosphatidylethanolamine is generated by PSD1. The spatially distinct pools within the cell are functionally distinct, but both are essential to life. For example, disruption of the phosphatidylserine decarboxylase gene causes misshapen mitochondria and has lethal consequences in embryonic mice, although phosphatidylethanolamine synthesis continues for a time in other cellular regions. Similarly, elimination of the endoplasmic reticulum route is embryonically lethal. Three additional minor biosynthetic pathways are known. Phosphatidylethanolamine can be formed by the enzymatic exchange reaction of ethanolamine with phosphatidylserine, or by re-acylation of lysophosphatidylethanolamine. The second of these is associated with the mitochondria-associated membrane where the phosphatidylserine synthase II is located. Finally, the bacterial plant pathogen Xanthomonas campestris is able to synthesize phosphatidylethanolamine by condensation of cytidine diphosphate diacylglycerol with ethanolamine. Ether lipids: It should be noted that all of these pathways for the biosynthesis of diacyl-phosphatidylethanolamine are very different and are separated spatially from that producing alkyl,acyl- and alkenyl,acyl-phosphatidylethanolamines, suggesting that there may be functional differences. In the protozoon T. brucei, for example, it has been demonstrated that the diacyl and ether pools of phosphatidylethanolamine have separate functions and cannot substitute for each other. Lands’ cycle: The various mechanisms produce different pools of phosphatidylethanolamine species, which are often in different cellular compartments and have distinctive compositions. Studies with mammalian cell types in vitro suggest that the CDP-ethanolamine pathway produces molecular species with mono- or di-unsaturated fatty acids on the sn-2 position preferentially, while the phosphatidylserine decarboxylation reaction generates species with polyunsaturated fatty acids on the sn-2 position mainly. However, as with other phospholipids, the final fatty acid composition in animal tissues is attained by a process of remodeling known as the Lands’ cycle . The first step is hydrolysis by a phospholipase A2 to lysophosphatidylethanolamine, followed by reacylation by means of various acyl-CoA:lysophospholipid acyltransferases. The enzymes LPCAT1, 2 and 3, which are involved in phosphatidylcholine biosynthesis, are also active with phosphatidylethanolamine, while LPEAT1 utilizes lysophosphatidylethanolamine mainly and is specific for oleoyl-CoA. Some of these isoforms appear to be confined to particular tissues. There is also a CoA-independent acyltransferase in macrophages that transfers arachidonic acid from phosphatidylcholine to ethanolamine-containing phospholipids. Phosphatidylethanolamine – Biological Function Physical properties: Although phosphatidylethanolamine has sometimes been equated with phosphatidylcholine in biological systems, there are significant differences in the chemistry and physical properties of these lipids, and they have different functions in biochemical processes. Both are key components of membrane bilayers and the ratio of the two may be important to many cellular functions. However, phosphatidylethanolamine has a smaller head group, which gives the lipid a cone shape. On its own, it does not form bilayers but inverted hexagonal phases. With other lipids in a bilayer, it is believed to exert a lateral pressure that modulates membrane curvature and stabilizes membrane proteins in their optimum conformations. It can hydrogen bond to adjacent lipids and to proteins through its polar head group. For example, the phosphate can be stabilized at the binding site by interactions with lysine and arginine side chains, or with hydroxyls from tyrosine side chains. There can also be strong interactions with acyl chains of the phospholipid. In contrast to phosphatidylcholine, phosphatidylethanolamine is concentrated with phosphatidylserine in the inner leaflet of the plasma membrane. On the other hand, it is present in both the inner and outer membranes of mitochondria, but especially the former, where it is present at much higher levels than in other organelles and facilitates membrane fusion and protein movement across membranes in addition to many other essential mitochondrial functions, including oxidative phosphorylation via the electron transport chain. It is an important functional component of membrane contact sites between the endoplasmic reticulum and mitochondria. In eukaryotic cells, especially those of insects, studies suggest that it has a similar function to cholesterol in membranes in that it increases the rigidity of the bilayer to maintain membrane fluidity. In bacterial membranes, it appears that a primary role for phosphatidylethanolamine is simply to dilute the high negative charge density of the anionic phospholipids. Protein Interactions: Membrane proteins amount to 30% of the genome, and they carry out innumerable biochemical functions, including transport, energy production, biosynthesis, signaling and communication. Within a membrane, most integral proteins consist of hydrophobic α-helical trans-membrane domains that zigzag across it and are connected by hydrophilic loops. Of those parts of the proteins outwith the bilayer, positively charged residues are much more abundant on the cytoplasmic side of membrane proteins as compared to the trans side (the positive-inside rule). Phosphatidylethanolamine is believed to have a key function in that it inhibits location of negative amino acids on the cytoplasmic side, supporting the positive-inside rule, and it has an appropriate charge density to balance that of the membrane surface and the protein. However, it can also permit the presence of negatively charged residues on the cytosolic surface in some circumstances in support of protein function. Phosphatidylethanolamine is vital for mitochondrial functionality, as demonstrated by defects in the operation of oxidative phosphorylation in the absence of PSD1, and by the role of the lipid in the regulation of mitochondrial dynamics in general and the biogenesis of proteins in the outer mitochondrial membrane. Synthesis of phosphatidylethanolamine in the inner membrane of mitochondria is critical for the function of the cytochrome bc1 complex (III), where there is a conserved binding site for the lipid in a specific subunit. Phosphatidylethanolamine binds non-covalently to a superfamily of cytosolic proteins with multiple functions termed 'phosphatidylethanolamine-binding proteins'. While four members have been identified in mammals (PEBP1-4), more than 400 members of the family that are conserved during evolution are known from bacteria to higher eukaryotes. These have many different functions including lipid binding, neuronal development, serine protease inhibition, and the regulation of several signaling pathways; in plants, they control shoot growth and flowering. PEBP4 is of particular interest as it is highly expressed in many different cancers and can increase their resistance to therapy. In animal tissues, phosphatidylethanolamine is especially important in the sarcolemmal membranes of the heart during ischemia, it is involved in the secretion of the nascent very-low-density lipoproteins from the liver and it has functions in membrane fusion and fission. It has a functional role in the Ca2+-ATPase in that one molecule of phosphatidylethanolamine is bound in a cavity between two transmembrane helices, acting as a wedge to keep them apart. This is displaced when Ca2+ is bound to the enzyme. Many other important proteins bind non-covalently to phosphatidylethanolamine in a similar way, including rhodopsin and aquaporins. The content of phosphatidylethanolamine in newly secreted VLDL particles and in apoB-containing particles isolated from the lumen of the Golgi is much higher than that in circulating VLDLs, suggesting that this lipid is involved in VLDL assembly and/or secretion. However, it is rapidly and efficiently removed from the VLDL in the circulation. With lipid droplets in cells, phosphatidylethanolamine is believed to promote coalescence of smaller droplets into larger ones. Although the mechanism has yet to be fully elucidated, effects on protein conformation are believed to be behind a finding that phosphatidylethanolamine is the primary factor in the brain required for the propagation and infectivity of mammalian prions. Host defense peptides are antimicrobial agents produced by both prokaryotic and eukaryotic organisms, and many of these have a high affinity for phosphatidylethanolamine as a lipid receptor to modulate their activities. For example, the peptide antibiotics cinnamycin and duramycins from Streptomyces have a hydrophobic pocket that fits around phosphatidylethanolamine such that the binding is stabilized by ionic interaction between the ethanolamine group of the lipid and the carboxylate moiety of the peptide; this complex exhibits activity against other Gram-positive organisms, such as Bacillus species. Much of the evidence for the unique properties of phosphatidylethanolamine comes from studies of the biochemistry of the bacterium E. coli, where this lipid is a major component of the membranes. Gram-negative bacteria have two membrane bilayers in the cell wall, and as much as 90% of the phospholipid in the inner leaflet of the outer membrane is phosphatidylethanolamine, with a high proportion in the cytoplasmic leaflet of the inner membrane also. In particular, phosphatidylethanolamine has a specific involvement in supporting the active transport of lactose by the lactose permease, and other transport systems may require or be stimulated by it. There is evidence that phosphatidylethanolamine acts as a 'chaperone' during the assembly of this and other membrane proteins to guide the folding path for the proteins and to aid in the transition from the cytoplasmic to the membrane environment, although in contrast it inhibits the folding of some multi-helical proteins. In the absence of this lipid, the transport membranes may not have the correct tertiary structure and so will not function correctly. Whether the lipid is required once the protein is correctly assembled is not fully understood in all cases, but it may be needed to orient enzymes correctly in the inner membrane. Some studies suggest that life in this organism can be maintained without phosphatidylethanolamine, but that life processes are inhibited. Autophagy and ferroptosis: A covalent conjugate of phosphatidylethanolamine with a protein designated 'Atg8' is formed by the action of cysteine protease ATG4 (belonging to the caspase family) and various other proteins, and is involved in the process of autophagy (controlled degradation of cellular components) in yeast by promoting the formation of membrane vesicles containing the components to be degraded (phosphatidylinositol 3-phosphate is also essential to this process). Similarly, oxidatively modified phosphatidylethanolamine is an important factor in ferroptosis, a form of apoptosis in which disturbances to iron metabolism lead to an accumulation of hydroperoxides. Precursor of other lipids: Phosphatidylethanolamine is a precursor for the synthesis of N-acyl-phosphatidylethanolamine (see below) and thence of anandamide (N‑arachidonoylethanolamine), and it is the donor of ethanolamine phosphate during the synthesis of the glycosylphosphatidylinositol anchors that attach many signaling proteins to the surface of the plasma membrane. In bacteria, it functions similarly in the biosynthesis of lipid A and other lipopolysaccharides. It is also the substrate for the hepatic enzyme phosphatidylethanolamine N-methyltransferase, which provides about a third of the phosphatidylcholine in the liver. Miscellaneous other functions: Phosphatidylethanolamine is the precursor of an ethanolamine phosphoglycerol moiety bound to two conserved glutamate residues in eukaryotic elongation factor 1A, which is an essential component in protein synthesis. This unique modification appears to be of great importance for the resistance of plants to attack by pathogens. Francisella tularensis bacteria, the cause of tularemia, suppresses host inflammation and the immune response when infecting mouse cells. The effect is due to a distinctive phosphatidylethanolamine species containing 10:0 and 24:0 fatty acids, and the synthetic lipid produces the same effects in vitro in human cells infected with dengue fever virus. It is hoped that this lipid will prove to be a potent anti-inflammatory therapeutic agent. Plants: In the seeds of higher plants, a deficiency of phosphorylethanolamine cytidylyltransferase, a rate-limiting enzyme in the biosynthesis of phosphatidylethanolamine, has profound effects on the viability and maturation of embryos. Lysophosphatidylethanolamine Figure \(22\) shows the the structure of lysophosphatidyethanolamine Lysophosphatidylethanolamine (LPE), with one mole of fatty acid per mole of lipid, is found in trace amounts only in animal tissues, other than plasma (10 to 50µM, or ~1% of total serum phospholipids). It is formed by hydrolysis of phosphatidylethanolamine by the enzyme phospholipase A2, as part of a de-acylation/re-acylation cycle that controls its overall molecular species composition as discussed above. A membrane-bound O-acyltransferase (MBOAT2) specific for LPE (and lysophosphatidic acid) has been characterized with a preference for oleoyl-CoA as substrate. There are reports of the involvement of LPE in cellular functions, such as differentiation and migration of certain neuronal cells, but also of various cancer cells. For example, oleoyl-LPE in the brain stimulates neurite outgrowth and protects against glutamate toxicity. In plants, lysophosphatidylethanolamine is a specific inhibitor of phospholipase D, a key enzyme in the degradation of membrane phospholipids during the early stages of plant senescence. Through this action, it retards the senescence of leaves, flowers, and post-harvest fruits. Indeed, it has a number of horticultural applications when applied externally, e.g., to stimulate ripening and extend the shelf-life of fruit, delay senescence, and increase the vase life of cut flowers. In bacteria, lysophosphatidylethanolamine displays chaperone-like properties, promoting the functional folding of citrate synthase and other enzymes. Some biological properties have been reported in animal tissues in vitro, but a specific receptor has yet to be identified. Lysophospholipids and especially lysophosphatidylethanolamines are produced in the envelope membranes of bacteria by many different endogenous and exogenous factors and must be transported back into the bacterial cell by flippases for conversion back to the diacyl forms by the action of a peripheral enzyme, acyl-ACP synthetase/LPL acyltransferase. Lysophosphatidylethanolamines produced by certain bacteria act synergistically with the sulfonolipid rosette-inducing factors (RIFs) to maximize the activity of the latter to induce choanoflagellates to move from a unicellular to a multicellular state. N-Acyl Phosphatidylethanolamine In N-Acyl phosphatidylethanolamine, the free amino group of phosphatidylethanolamine is acylated by a further fatty acid. This lipid has been detected in rather small amounts in several animal tissues (~0.01%), but especially brain, nervous tissues, and the epidermis, when the N-acyl chain is often palmitic or stearic acid (human plasma: N16:0-PE (40%), N18:1-PE (23.3%), N18:0-PE (19%), N18:2-PE (16.6%) and N20:4-PE (1.4%)). Under conditions of degenerative stress, it can accumulate in significant amounts, for example as the result of ischemic injury, infarction, or cancer. It is present in plasma after feeding a high-fat diet to rats, and then it can cross into the brain where it accumulates in the hypothalamus. Figure \(23\) shows the structures of N-Acyl Phosphatidylethanolamine. In animals, N-Acyl phosphatidylethanolamine is of particular importance as the precursor of anandamide, and of other biologically important ethanolamides (e.g., N-oleoylethanolamide) in brain and other tissues, but especially the intestines. In brief, it is formed biosynthetically by the action of a transferase (cytosolic phospholipase A2ε) exchanging a fatty acid from the sn-1 position of a phospholipid (probably phosphatidylcholine) to the primary amine group of phosphatidylethanolamine (without a hydrolytic step). Both diacyl- and alkenylacyl-species of phosphatidylethanolamine can serve as acceptors. In addition, some transfer can also occur from phosphatidylethanolamine per se by an intramolecular reaction. However, it should be noted that some N-acyl phosphatidylethanolamine can be formed artefactually as a result of faulty extraction procedures during analysis. In plants, N-acyl phosphatidylethanolamine is a common constituent of cereal grains (e.g., wheat, barley and oats) and of some other seeds (1.9% of the phospholipids of cotton seeds, but 10-12% of oats). In other plant tissues, it is detected most often under conditions of physiological stress. In contrast to animals, synthesis involves direct acylation of phosphatidylethanolamine with a free fatty acid, catalyzed by a membrane-bound transferase in a reverse serine-hydrolase catalytic mechanism. Activation of N-acyl phosphatidylethanolamine metabolism in plants with the release of N-acylethanolamines and phosphatidic acid formation seems to be associated with cellular stresses, but research is at an early stage. However, both N-acyl lipid classes have been implicated in such physiological processes as the elongation of main and lateral roots, regulation of seed germination, seedling growth, and defense from attacks by pathogens. N-Acyl phosphatidylethanolamine has been found in a number of microbial species, while N-acetyl phosphatidylethanolamine was detected in a filamentous fungus, Absidia corymbifera, where it comprised 6% of the total membrane lipids. It was accompanied by an even more unusual lipid 1,2‑diacyl-sn-glycero-3-phospho(N-ethoxycarbonyl)-ethanolamine. Phosphatidylserine and Related Lipids Phosphatidylserine or 1,2-diacyl-sn-glycero-3-phospho-L-serine is an important anionic phospholipid, which brings essential physical properties to membranes in both eukaryotes and prokaryotes. Independently of this, it has many biological functions in cells, including effects on blood coagulation and apoptosis, and it is the biosynthetic precursor for phosphatidylethanolamine in prokaryotes and in eukaryote mitochondria. Its metabolite lysophosphatidylserine has signaling functions and operates through specific receptors. Also, there is increasing interest in a structurally related lipid phosphatidylthreonine, and other phospholipids linked to amino acids. Phosphatidylserine - Structure and Occurrence Although phosphatidylserine is distributed widely among animals, plants, and microorganisms, it is usually less than 10% of the total phospholipids, the greatest concentration being in myelin from brain tissue. For example, mouse brain and liver contain 14 and 3% phosphatidylserine, respectively. However, it may comprise 10 to 20 mol% of the total phospholipids in the plasma membrane, where under normal conditions it is concentrated in the inner leaflet, and in the endoplasmic reticulum of cells. In the yeast Saccharomyces cerevisiae, it is a minor component of most cellular organelles other than the plasma membrane, where surprisingly it can amount to more than 30% of the total lipids. In most bacteria, it is a minor membrane constituent, although it is important as an intermediate in phosphatidylethanolamine biosynthesis. The 1‑octadecanoyl-2-docosahexaenoyl molecular species, which is especially important in brain tissue, is illustrated here. Figure \(24\) shows the structure of phosphatidylserine Phosphatidylserine is an acidic (anionic) phospholipid with three ionizable groups, i.e., the phosphate moiety, the amino group, and the carboxyl function. As with other acidic lipids, it exists in nature in salt form, but it has a high propensity to chelate to calcium via the charged oxygen atoms of both the carboxyl and phosphate moieties, modifying the conformation of the polar head group. This interaction may be of considerable relevance to the biological function of phosphatidylserine, especially during bone formation for example. In animal cells, the fatty acid composition of phosphatidylserine varies from tissue to tissue, but it does not appear to resemble the precursor phospholipids, either because of selective utilization of specific molecular species for biosynthesis or because of the re-modeling of the lipid via deacylation-reacylation reactions with lysophosphatidylserine as an intermediate (see below). In human plasma, 1-stearoyl-2-oleoyl and 1-stearoyl-2-arachidonoyl species predominate, but in the brain (especially grey matter), retina and many other tissues 1-stearoyl-2-docosahexaenoyl species are especially abundant and appear to be essential for normal functioning of the nervous system. Indeed, the ratio of n-3 to n-6 fatty acids in brain phosphatidylserine is much higher than in most other lipids. The positional distribution of fatty acids in phosphatidylserine from rat liver and bovine brain are listed in Table \(5\)​​​​​​​. As with most phospholipids, saturated fatty acids are concentrated in position sn-1 and polyunsaturated in position sn-2. Table \(5\)​​​​​​​: Positional distribution of fatty acids in phosphatidylserine from rat liver and bovine brain Position Fatty acid 16:0 18:0 18:1 18:2 20:4 22:6 Rat liver [1] sn-1 5 93 1 sn-2 6 29 8 4 32 19 Bovine brain [2] sn-1 3 81 13 sn-2 2 1 25 trace 1 60 1. Wood, R. and Harlow, R.D. Arch. Biochem. Biophys., 135, 272-281 (1969); DOI. 2. Yabuuchi, H. and O'Brien, J.S. J. Lipid Res., 9, 65-67 (1968); DOI. In leaves of Arabidopsis thaliana, used as a 'model' plant in many studies, the fatty acid composition of phosphatidylserine resembles that of phosphatidylethanolamine. There is an intriguing report that the chain lengths of the acyl groups increase with age and stress in phosphatidylserine quite specifically, and 22:0 and 24:0 fatty acids have been reported to occur in this lipid in the plasma membrane of some plant species. In marked contrast to phosphatidylethanolamine, phosphatidylserines with ether-linked moieties (alkyl and alkenyl) are not common in animal tissues, although they are reported to be relatively abundant in human retina and macrophages (they were first found in rat lung). As a generality, the concentration of phosphatidylserine is highest in plasma membranes and endosomes but is very low in mitochondria. As it is located entirely on the inner monolayer surface of the plasma membrane (and of other cellular membranes) and it is the most abundant anionic phospholipid, it may make the largest contribution to interfacial effects in membranes involving non-specific electrostatic interactions. This normal distribution is disturbed during platelet activation and cellular apoptosis. N-Acylphosphatidylserine is reportedly present in the frontal cortex of patients with schizophrenia, as a minor component of the lipids of sheep erythrocytes, bovine brain, and the central nervous system of freshwater fish, and Bryozoans amongst others. The N-arachidonoyl form may be the precursor of the endocannabinoid N-arachidonoylserine. Biosynthesis of Phosphatidylserine L-Serine is a non-essential amino acid that is actively synthesized by most organisms. In animals, it is produced in nearly all cell types, although in brain it is synthesized by astrocytes but not by neurons, which must be supplied with this amino acid for the biosynthesis of phosphatidylserine (and of sphingoid bases). In animal tissues, phosphatidylserine is synthesized solely by calcium-dependent base-exchange reactions in which the polar head-group of an existing phospholipid is exchanged for L-serine. There are two routes involving distinct enzymes (PS synthase I and II) with 30% homology and several membrane-spanning domains that can utilize different substrates. Phosphatidylserine is synthesized by both enzymes on the cytosolic face of the endoplasmic reticulum (ER) of the cell, but mainly in a specific domain of this termed the mitochondria-associated membrane ('MAM'), because it is tethered transiently to the mitochondrial outer membrane, presumably by a protein bridge. In yeast, a complex of integrated proteins ('ERMES') has been characterized with a similar function. The reaction involves the exchange of L-serine with either phosphatidylcholine or phosphatidylethanolamine, catalyzed by PS synthase I (although it was long thought that only phosphatidylcholine was a substrate for this enzyme), while PS synthase II catalyzes a similar exchange with diacyl-phosphatidylethanolamine and its the plasmalogen form. Both enzymes are subject to feedback regulation by their product phosphatidylserine, thereby maintaining the correct amounts of this lipid. Figure \(25\) shows the synthesis and metabolism of phosphatidylserine in animals. Phosphatidylserine synthase I is expressed in all mouse tissues, but especially the kidney, liver, and brain, while phosphatidylserine synthase II is most active in the brain and testis and much less so in other tissues. The latter enzyme has a high specificity for molecular species containing docosahexaenoic acid. It is not known why such a complex series of coupled reactions is necessary, or why there should be two enzymes, but one virtue is that the free ethanolamine and choline formed are rapidly re-utilized for phospholipid synthesis. Thus, both phosphatidylserine and phosphatidylethanolamine are produced without a reduction in the amount of phosphatidylcholine. Elimination of both enzymes is embryonically lethal in knock-out mice, but each of them can be knocked out separately and the mice survive, even though they have substantially reduced levels of phosphatidylserine and phosphatidylethanolamine. As with other phospholipids, the final fatty acid composition in animal tissues is attained by a process of remodeling known as the Lands’ cycle. The first step is hydrolysis by a phospholipase A2 to lysophosphatidylserine, followed by the reacylation by various acyl-CoA:lysophospholipid acyltransferases. One membrane-bound O-acyltransferase (LPCAT4 or MBOAT2) with a preference for oleoyl-CoA has been characterized, while a second (LPCAT3 or MBOAT5) incorporates linoleoyl and arachidonoyl chains (and also utilizes lysophosphatidylcholine). Following synthesis, phosphatidylserine molecules can diffuse laterally in a concentration-dependent manner to different regions of the membrane to fulfill their physiological functions. In humans, cytosolic transport proteins transfer phosphatidylserine and other acidic phospholipids between membranes, and this can also occur by a vesicular transport mechanism. Some of the newly synthesized phosphatidylserine is transferred to the plasma membrane, while a proportion is transported to the mitochondria, probably again via transient membrane contact (MAM), where it is decarboxylated to produce phosphatidylethanolamine by a specific decarboxylase in the inner mitochondrial membrane. In yeast, there is a preference for molecular species containing two monoenoic fatty acids for transport and metabolism; this process occurs also at the Golgi/endosome membranes. All the phosphatidylethanolamine in mitochondria is formed in this way, but some can return to the endoplasmic reticulum where it may be converted back to phosphatidylserine by the action of the PS synthases. Mitochondrial production of phosphatidylethanolamine from phosphatidylserine is not fully complemented by the CDP-ethanolamine pathway, as mice lacking the enzyme do not survive for long. Evidently, cellular concentrations of these two lipids are intimately related and tightly regulated. Figure \(26\) shows the mitochondrial metabolism of phosphatidylserine Much of the phosphatidylserine thus formed is decarboxylated to phosphatidylethanolamine, and this may be the major route to the latter in bacteria. As phosphatidylcholine in yeast is produced via methylation of phosphatidylethanolamine, phosphatidylserine is the primary precursor for this phospholipid in these organisms. Bacteria and plants: In bacteria and other prokaryotic organisms and in yeast, phosphatidylserine is synthesized by a mechanism comparable to that of most other phospholipids, i.e., by reaction of L-serine with CDP-diacylglycerol, and depends on Mg2+ or Mn2+. Phosphatidylserine synthases belong to two different families: type I (non-integral membrane form) in the phospholipase D-like family as in E. coli, and type II (integral membrane form) in the CDP-alcohol phosphotransferase family as in Bacillus sp. and the yeast S. cerevisiae, although the latter shows no homology with the bacterial enzymes. In many plants, including in the model plant Arabidopsis, much of the phosphatidylserine is produced by a calcium-dependent base-exchange reaction in which the head group of an existing phospholipid is exchanged for L-serine in the luminal leaflet of the endoplasmic reticulum (i.e., mechanistically similar to PS synthase I). It is transferred to the cytoplasmic membrane leaflet by flippases and thence to the post-Golgi compartments before eventually accumulating at the plasma membrane. However, some vesicular transport may occur or there may be direct transfer at membrane contact sites. A CDP-diacylglycerol (prokaryotic-like) biosynthetic pathway exists in some species, e.g. wheat. Let's explore the mechanism of the Methanocaldococcus jannaschii phosphatidylserine synthase (MjPSS). The organism is a hyperthermophilic methanogen. Figure \(27\) shows substrate binding by MjPSS. The large binding pocket for CDP-DAG in MjPSS extends from the hydrophobic membrane core to the active site near the cytoplasmic surface in the center of the dimer. In the closed structures (left), both CDP-DAG alkyl chains adopt similar conformations within the binding pocket, whereas the positions of helix 7 and 8 in the open structures (right) allow one alkyl chain to reach the membrane via a different path. Serine molecules are only found in open structures (right). In one closed conformation, there is a citrate near the substrate-binding site, whereas in the other closed structures this position is empty. A chain of three chloride ions extends parallel to the dimer interface from the active site to the cytoplasmic interface with the N-terminal helix hH from the other protomer. Figure \(28\) shows the reaction cycle of MjPSS during the synthesis of PS from CDP-DAG and serine in the presence of Mg2+ and Ca2+. The CDPDAG binding site in MjPSS of is formed and stabilized by the divalent cations Mg2+ and Ca2+ (a). In the absence of CDP-DAG, Ca2+ most likely is coordinated by water molecules, as shown in a, or by residues in nearby loops that would be flexible in the absence of CDP-DAG. The binding of CDP-DAG (b) is driven by the coordination of Ca2+ by the negatively charged phosphates. Serine binds to the binding pocket after CDP-DAG (c). For the nucleophilic attack, the serine molecule is positioned with its hydroxyl group near the β-phosphate of CDP-DAG. The serine molecule is activated by deprotonation, attacks the β-phosphate, and forms the penta-coordinated transition state (d). The proton of serine is probably removed by one of the water molecules located in the interaction network of Asp66, Arg101, and the nearby chloride ions. Hydrolysis of the CDP-DAG/serine complex from the transition state leads to the complex of MjPSS with the products PS and CMP (e). The next cycle starts after release of the products and binding of CDP-DAG. Structural data are available for the state with bound CDP-DAG (b), CDP-DAG, and serine (c), and for the transition state of the CDP-DAG/serine complex (d). Figure \(29\) shows an interactive iCn3D model of the Methanocaldococcus jannaschii phosphatidyl serine synthase (PSS) in the open state with bound CDP-DAG and serine (7B1L). The enzyme is also named CDP-diacylglycerol--serine O-phosphatidyltransferase. Figure \(29\): Bacterial phosphatidyl serine synthase (PSS) in the open state with bound CDP-DAG and serine (7B1L) (Note the actual PDB file title names state that this is the closed state which it is not.) Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...B3Rn3gVN4c4ge6 The A chain of the homodimer is shown in gray and the B in plum. The active site residues in the A chain in the above mechanism are shown in stick, CPK colors, and labeled. Hover over the large ligand (58A) in the gray subunit. Two free serines are shown near it but only one is probably the substrate. Phosphatidylserine – Biological Function Membrane location: Phosphatidylserine modulates membrane charge locally, enabling the recruitment of soluble cations and proteins, and so it contributes to the organization of processes within cell membranes. Its distribution within membranes is tightly controlled as it facilitates signaling within the various cellular compartments. Thus, it undergoes a transition from the lumenal leaflet of the endoplasmic reticulum to the cytosolic leaflet in the trans-Golgi network, probably by the activity of flippases and scramblases in the Golgi, and it is highly enriched on the inner leaflet of the plasma membrane. Transport to the plasma membrane against a concentration gradient is aided in part by proteins designated 'ORP5' and 'ORP8' in humans (Osh 6 and Osh7 in yeast) with a 'PH' binding domain for phosphatidylinositol 4,5-bisphosphate and an 'ORD' domain for phosphatidylserine. At a membrane contact site between the endoplasmic reticulum and plasma membrane, phosphatidylserine is exchanged for phosphatidylinositol 4-phosphate. Such transfer requires an input of energy, which can be supplied in the form of ATP or by phosphoinositides Although it does not take part in membrane raft formation, phosphatidylserine is present in caveolae, where it is believed to interact with caveolin-1. It is also present in appreciable amounts in the endosomal compartment. The asymmetric structure of the plasma membrane with high concentrations of anionic lipids such as phosphatidylserine in the cytosolic leaflet with zwitterionic lipids in the extracellular leaflet generates two surfaces with greatly different electrostatic potentials that influence the association of proteins with the membrane surface and the activities of integral membrane proteins. This distribution is maintained and can be altered, after specific activation, by various flippases (transfer back into the cytoplasmic leaflet), floppases (transfer out of the cytoplasmic leaflet), and scramblases (bidirectional transfer), including ATP-dependent translocases selective for phosphatidylserine. Phosphatidylserine is highly enriched in the cytosolic leaflet of the membranes of recycling endosomes, which replenish the lipids and proteins of the plasma membrane, and it is essential for their function. Enzyme activation: In addition to its function as a component of cellular membranes and as a precursor for other phospholipids, phosphatidylserine is an essential cofactor that binds to and activates a large number of proteins, especially those with signaling activities. The negative charge on the lipid facilitates the binding to proteins through electrostatic interactions or Ca2+ bridges. For example, the presence of appreciable amounts of phosphatidylserine on the cytosolic leaflet of endosomes and lysosomes enables these compartments to dock with proteins that possess specific phosphatidylserine-binding domains including several important signaling and fusogenic effectors. The cytoskeletal protein spectrin binds to phosphatidylserine in this way, and it is also required by enzymes such as the neutral sphingomyelinase and the Na+/K+ ATPase, where the 18:0/18:1 molecular species is especially important. It is believed that the fatty acyl components of this species in the inner leaflet of the plasma membrane (and potentially other intracellular membranes) may interact (interdigitation or "hand-shake") with the very-long chains of sphingolipids in the outer leaflet in raft microdomains, resulting in a high local concentration of the anionic phospholipid and an accumulation of negative surface charge to which specific poly-cationic proteins in the membranes can bind. This may then enable the transfer of signals across the membrane to the cytosol. Similarly, phosphatidylserine participates directly in key signaling pathways in the brain by binding to the cytosolic proteins involved in neuronal signaling and thereby activating them. At least three major pathways are affected, including those involving phosphatidylinositol 3-kinase and protein kinase C. For example, most enzymes of the protein kinase C family contain a 'C2' calcium-dependent cysteine-rich region that recognizes phosphatidylserine, and in coordination with the 'C1' domain that binds to diacylglycerols, is essential for activating and locating them to the plasma membrane of appropriately stimulated cells. Phosphatidylserine is not involved in cell signaling through the formation of metabolites, as is the case with phosphatidylinositol. Blood coagulation: Phosphatidylserine is an important element of the blood coagulation process in platelets, where it is transported from the inner to the outer surface of the plasma membrane in platelets activated by exposure to fibrin-binding receptors, for example. Here, the exposed phosphatidylserine enhances the activation of prothrombin to thrombin (the key molecule in the blood clotting cascade) by triggering a cascade of reactions and providing the negatively charged platform that enables calcium ions to form bridges with γ-carboxyglutamic acid-containing domains on the coagulation factors. Membrane vesicles with phosphatidylserine exposed on the surface can also be released from platelets and promote the coagulation process. Apolipoprotein A-1 in high-density lipoproteins has a controlling function in that it neutralizes these procoagulant properties by arranging the phospholipid in surface areas that are too small to accommodate the prothrombinase complex. Blood coagulation is beneficial when it prevents the loss of blood from the circulatory system, but it is detrimental when it causes thrombosis, and the action of phosphatidylserine is essential to the regulation of the process. Apoptosis: In addition in response to particular calcium-dependent stimuli, phosphatidylserine is known to have an important role in the regulation of apoptosis or programmed cell death, the natural process by which aged or damaged cells are removed from tissues before they can exert harmful effects. When cells are damaged, a mechanism is initiated in which the normal distribution of this lipid on the inner leaflet of the plasma membrane bilayer is disrupted by stimulation of scramblases, which are ATP-independent and can move the lipid across the membrane to the outer leaflet. This occurs together with the inhibition of aminophospholipid translocases, which return the lipid to the inner side of the membrane. In erythrocytes, phosphatidylserine is located in the inner leaflet of the membrane bilayer under low Ca2+ conditions when a phospholipid scramblase is suppressed by membrane cholesterol, but it is exposed to the outer leaflet under elevated Ca2+ concentrations which activate the scramblase. After the collapse of this asymmetry and transfer of phosphatidylserine to the outer leaflet of an effete cell, it is believed that it is recognized by a cohort of receptors, either directly or indirectly, through bridging ligands on the surface of macrophages and related scavenger cells. These activate a family of cysteine-dependent aspartate-specific proteases, the caspases, and other enzymes to facilitate the engulfment of the apoptotic cells and their potentially toxic or immunogenic contents in a non-inflammatory manner. It is noteworthy that the transition from a pro-inflammatory to an anti-inflammatory state is defined by phagocytosis of neutrophils by macrophages via this phosphatidylserine-dependent process. During apoptosis, the generation of reactive oxygen species occurs, mainly hydrogen peroxide, which together with the enzyme cytochrome c brings about rapid oxidation of the fatty acids in phosphatidylserine before this lipid is externalized. Indeed, it is now apparent that molecular species of phosphatidylserine with an oxidatively truncated sn-2 acyl group that incorporates terminal γ-hydroxy(or oxo)-α,β-unsaturated acyl moieties are especially potent signals for scavenger receptors in macrophages as a prerequisite for engulfment of apoptotic cells. This has been described as "a dominant and evolutionarily conserved immunosuppressive signal that promotes tolerance and prevents local and systemic immune activation" or more succinctly as an "eat-me signal" (externalized phosphatidylinositol 3,4,5-trisphosphate (PI(3,4,5)P) may have a similar function). The binding of phosphatidylserine to specific proteins, such as apolipoprotein H (β2-glycoprotein 1), enhances the recognition and clearance. This process is essential for the development of the lung and brain, and it is also relevant to clinical situations where apoptosis plays an important part, such as cancer, chronic autoimmunity, and infections. For example, phosphatidylserine is a necessary component of the TAM family of receptor tyrosine kinases and the receptor-ligand complex of particular importance in cancer cells, where phosphatidylserine-TAM signaling regulates many aspects of inflammation and immune resolution and is seen as a target for therapeutic intervention. Exposure of phosphatidylserine is increased substantially on the surface of tumor cells or tumor cell-derived microvesicles, which have innate immunosuppressive properties and facilitate tumor growth and metastasis. Targeting phosphatidylserine is considered to be a promising strategy in cancer immunotherapy. In relation to atherosclerosis, phosphatidylserine is believed to have anti-inflammatory and protective effects as a component of the high-density lipoproteins, probably mediated by the apoptosis mechanism. In contrast, as this mechanism is important for the turnover of erythrocytes, it is relevant to thrombus formation and the stabilization of blood clots. The innate immunosuppressive effect of externalized phosphatidylserine has been hijacked by numerous viruses and bacteria to facilitate infection. A similar apoptotic mechanism operates in retinal pigment epithelial cells to remove the large amounts of photoreceptor cell debris that are generated continuously. In addition, appreciable amounts of phosphatidylserine are translocated by an analogous mechanism to the surface of T lymphocytes that express low levels of the trans-membrane enzyme tyrosine phosphatase. This change in distribution acts then as a signaling mechanism to modulate the activities of several membrane proteins. The anti-coagulant protein annexin V binds with high specificity to phosphatidylserine and is used as a probe to detect apoptotic cells. It is noteworthy that phosphatidylserine is a major component of the membranes of microvesicles in animal cells, and translocation to the outer leaflet upon cellular activation is essential for their biogenesis. In addition, exposure of phosphatidylserine on the cell surface is reported to be a factor in non-apoptotic forms of regulated inflammatory cell death, such as necroptosis. Role in infections: Unfortunately, viruses such as Ebola and HIV viruses can hijack this apoptosis machinery by incorporating phosphatidylserine into their viral envelopes so conning cells into engulfing them; the viral glycoprotein/cellular receptor complex may then facilitate the entry of foreign organisms into other cells. Similarly, parasites ingested in this manner, including Leishmania and Trypanosoma species, utilize host phosphatidylserine to establish infections and facilitate disease progression as they do not then elicit the production of proinflammatory cytokines. This mechanism has been termed 'apoptotic mimicry' and is critical for the survival of parasites within the macrophage. Other activities: Phosphatidylserine is required for the transmembrane movement of excess cholesterol, derived initially from the lysosomal degradation of low-density lipoproteins, from the plasma membrane to the endoplasmic reticulum thereby maintaining membrane integrity and ensuring cell survival. It is therefore an important element in cholesterol homeostasis. The mechanism is believed to involve proteins known as GRAMD1s embedded in the endoplasmic reticulum membrane at sites in contact with the plasma membrane. These have two functional domains: the StART-like domain that binds cholesterol and the GRAM domain that binds anionic lipids, such as phosphatidylserine, and so forms a link between the two membranes that enables the transfer of cholesterol. A further unusual function of phosphatidylserine is that it is a key component of the lipid-calcium-phosphate complexes that act as nucleation centers for hydroxyapatite formation and initiate mineral deposition during the formation of bone. It has been established that phosphatidylserine and inorganic phosphate must be present, before calcium ions are introduced, when the high affinity of phosphatidylserine for calcium ions becomes important. Nucleation is facilitated by the protein annexin V. Similarly, during bone repair and maintenance, the fusion of osteoclasts requires the non-apoptotic exposure of phosphatidylserine at the surface of fusion-committed cells with the aid of a transmembrane protein (DC-STAMP) expressed in dendrocytes. This activity is relevant to cardiovascular disease and in particular to the phenomenon of "hardening of the arteries," where atherosclerotic plaques can undergo mineralization with the deposition of hydroxyapatite. Among many other functions of phosphatidylserine, it is believed to be an essential surface membrane component for the fusion of cell types other than osteoclasts, including during the formation of fibers in muscle cells, and the fusion of macrophages into inflammatory giant cells and myoblasts into myotubes. Such cell fusions require the non-apoptotic exposure of phosphatidylserine at the surface of fusing cells, where it interacts with phosphatidylserine-recognizing proteins to regulate the time and place of cell fusion. Phosphatidylserine provides stable membrane domains in spermatozoa that are essential for fertilization, and it is also an essential component of the plasma membrane microdomains known as caveolae, where it is required both for their formation and stability possibly through specific binding to the cavin proteins. The high concentrations of docosahexaenoic acid (DHA) in the brain and retinal phosphatidylserine are certainly important for the development and function of these tissues. Accumulation of phosphatidylserine in neuronal membranes is promoted by DHA, and this is important for the maintenance of neuronal survival. Phosphatidylserine may also be a reservoir of DHA for protectin formation in neuronal tissue. On the other hand, the Food and Drug Administration in the USA considers that there is little scientific evidence to support claims that dietary supplements of phosphatidylserine reduce the risk of dementia or cognitive dysfunction in the elderly, and other nutritional claims appear to be dubious also. Antibodies to phosphatidylserine are formed in some disease states, including thrombosis and recurrent spontaneous pregnancy loss. The rare genetic disease Lenz-Majewski syndrome is caused by a mutation in the phosphatidylserine synthase I gene that greatly increases the activity of the enzyme while preventing feedback inhibition, and abnormal metabolism of phosphatidylserine has been implicated in other diseases. In yeasts such as Candida albicans, phosphatidylserine and the enzyme phosphatidylserine decarboxylase, which generates phosphatidylethanolamine, are both essential for the virulence of the organism towards a host species. Lysophosphatidylserine Figure \(30\) shows the structure of lysophosphatidylserine Lysophosphatidylserine, i.e., with a fatty acid in one position only, is known to be a mediator of a number of biological processes, especially in the context of the immune system in animal tissues. It has been found in the thymus, peripheral lymphoid tissues, central nervous system, and colon, but is barely detectable in plasma. Deacylation of the diacyl lipid by phospholipases is the primary source. For example, a secreted isoform that is phosphatidylserine-specific (PLA1A) removes the sn-1 acyl group of phosphatidylserine to generate sn‑2‑lysophosphatidylserine containing unsaturated fatty acids, and this is upregulated greatly by various inflammatory stimuli. This extracellular enzyme utilizes phosphatidylserine exposed on the cell membrane as a substrate, although other phospholipases may operate intracellularly and produce sn‑1‑lysophosphatidylserine. In addition, platelets in some species (not significantly in humans) secrete a phospholipase A2 group IIA (ABHD16A), which generates saturated sn‑1‑lysophosphatidylserine (and other lysophospholipids). Lysophosphatidylserine has been detected after injury to animal tissues (tumor growth, graft rejection, burns), and it may have a similar function to lysophosphatidic acid in cell signaling, for example in regulating calcium flux and stimulating immune cells through G protein-coupled receptors of which three (GPR34, P2Y10 and GPR174, LPS1-3) have been detected in mice and humans. For example, GPR174 mediates the suppression of T-cell proliferation induced in vitro by lysophosphatidylserine. When cells are damaged, lysophosphatidylserine can be generated by a reaction dependent on the activation of the NADPH oxidase. It can diffuse and transmit the information to other cells, especially mast cells, and it is produced to enhance the clearance of activated and dying neutrophils. It thus has a role in the resolution of inflammation. One specific molecular species, i.e., 1‑(11Z‑eicosenoyl)-glycero-3-phosphoserine, is reported to be a true agonist of the Toll-like receptor 2/6 heterodimer of importance to the immune response to pathogens; both its polar head group and the length of the acyl chain are required for this activity. On the other hand, sn-2-lysophosphatidylserine has proinflammatory reactions in that it augments mast cell degranulation and mast cell-dependent anaphylactic shock; most other lysophospholipids have no such activity. Deregulated lysophosphatidylserine metabolism has been linked to certain cancers, cardio-metabolic disorders, night blindness, and the human genetic neurological disorder PHARC. High serum levels of PLA1A are associated with such autoimmune disorders as Graves' disease and systemic lupus erythematosus, and there is increased expression of the enzyme in metastatic melanomas. It is necessary for the assembly of the hepatitis C virus, and it can play a role in the antivirus innate immune response. In Schistosome infections, lysophosphatidylserine from the parasite is believed to be a key activator molecule in the host. Negatively charged lysophosphatidylserine species tend to organize in non-bilayer structures and are believed to facilitate the folding of certain membrane proteins in situ better than bilayer-forming lipids. Phosphatidylinositol and Related Phosphoinositides Although it had long been recognized that phosphatidylinositol or 1,2-diacyl-sn-glycero-3-phospho-(1'-myo-inositol) was a key membrane constituent, it was initially something of a surprise when the manifold biological activities of this lipid, and then of the derived phosphatidylinositol phosphates and their hydrolysis products, were discovered in animals, plants, and microorganisms. Many years after the initial discoveries in the 1950s, these lipids continue to be a major focus for research efforts around the world with considerable relevance to human health. Phosphatidylinositol and its various metabolites and relevant enzymes can be located and function within different membrane regions in cells, and they form part of what have been termed phosphoinositide and phosphatidylinositol cycles, their versatility stemming from the inositol head group, a six-carbon the hexahydroxy ring, which can be reversibly phosphorylated on the 3, 4 and 5 positions. In addition to their structural role in membranes, these lipids are intimately involved in innumerable aspects of membrane trafficking and signaling in eukaryotic cells, functions that are essential to cell growth and metabolism. Only a brief overview of such a highly complex topic is possible here. Glycosyl-phosphatidylinositol (GPI) is a related lipid that serves as an anchor for proteins. Phosphatidylinositol Structure and Occurrence: Phosphatidylinositol is an important lipid, both as a membrane constituent and as a participant in essential metabolic processes in all plants and animals, both directly and via a number of its metabolites. It is an acidic (anionic) phospholipid that in essence consists of a phosphatidic acid backbone, linked via the phosphate group to inositol (hexahydroxycyclohexane). In most organisms, the stereochemical form of the last is myo-D-inositol (with one axial hydroxyl in position 2 with the remainder equatorial, i.e. a chair-like structure), although other forms (scyllo- and chiro-) have been found on occasion in plants. The 1‑stearoyl,2-arachidonoyl molecular species, which is of considerable biological importance in animals, is illustrated. Figure \(31\) shows the structure of phosphatidylinositol. Phosphatidylinositol is especially abundant in brain tissue, where it can amount to 10% of the phospholipids, but it is present in all tissues, cell types, and membranes at relatively low levels in comparison to many other phospholipids. In rat liver, it amounts to 1.7 micromoles/g., i.e. less than phosphatidylcholine, phosphatidylethanolamine, and phosphatidylserine. Under normal conditions, it is present entirely in the inner leaflet of the erythrocyte membrane and of the plasma membrane in nucleated cells. Phosphatidylinositol per se is rarely found in prokaryotes other than the Actinomycetales, although the thermophilic α-proteobacterium Rhodothermus marinus contains dialkylether glycerophosphoinositides. The fatty acid composition of phosphatidylinositol is rather distinctive as shown in Table \(6\)​​​​​​​. Thus, in almost all animal tissues, the characteristic feature is a high content of stearic and arachidonic acids. All the stearic acid is linked to position sn-1 and all the arachidonic acid to position sn-2, and as much as 78% of the total lipid may consist of the single molecular species sn-1-stearoyl-sn-2-arachidonoyl-glycerophosphorylinositol (see Table \(7\)​​​​​​​​​​​​​​ below). Although 1-alkyl- and alkenyl- forms of phosphatidylinositol are known, they tend to be much less abundant than the diacyl form. In plant phosphatidylinositol, e.g. Arabidopsis thaliana as listed, palmitic acid is the main saturated fatty acid in position sn-1, while linoleic and linolenic acids are the main unsaturated components in position sn-2. Similarly in yeast, palmitic acid is in position sn-1 with oleic and palmitoleic acids in position sn-2 predominantly; the Amoebozoa have a C16 alkyl group in position sn-1 and cis-vaccenic acid in position sn-2. Table \(6\): ​​​​​​​ Fatty acid composition of phosphatidylinositol (wt % of the total) in animal and plant tissues. Tissue Fatty acids 16:0 18:0 18:1 18:2 18:3 20:3 20:4 22:3 22:5 22:6 Bovine brain [1] 8 38 10 1 - 5 34 2 tr. 1 Bovine liver [2] 5 32 12 6 1 7 23 4 3 5 Rat liver [3] 5 49 2 2   4 35     1 A. thaliana [4] 48 3 2 24 24 [1] = Holub, B.J. et al.. J. Lipid Res.., 11, 558-564 (1970); DOI. [2] = Thompson, W. and MacDonald, G., J. Biol. Chem., 250, 6779-6785 (1975); DOI. [3] = Wood, R. and Harlow, R.D. Arch. Biochem. Biophys., 135, 272-281 (1969); DOI. [4] = Browse, J. et al. Biochem. J., 235, 25-31 (1986); DOI. Biosynthesis: The basic mechanism for the biosynthesis of phosphatidylinositol and phosphatidylglycerol is sometimes termed a branch point in phospholipid synthesis, as phosphatidylcholine and phosphatidylethanolamine are produced by a somewhat different route. Phosphatidylinositol is found in all eukaryotes, which are in general able to synthesize inositol de novo via glucose-6-phosphate. As with phosphatidylglycerol (and hence cardiolipin), phosphatidylinositol is formed biosynthetically from phosphatidic acid via the intermediate cytidine diphosphate diacylglycerol, which is produced by the action of a CDP-diacylglycerol synthase believed to be the rate-limiting enzyme in phosphatidylinositol biosynthesis. Then, the enzyme CDP-diacylglycerol inositol phosphatidyltransferase ('phosphatidylinositol synthase' or 'PIS') catalyzes a reaction with myo-inositol to produce phosphatidylinositol. Figure \(32\) shows the synthesis of phosphatidylinositol in eukaryotes. Only isoform of PIS exists in mammals and it is located in the endoplasmic reticulum, in part in a subcompartment of this associated with mitochondria (mitochondria-associated membranes - MAM) and in mitochondria per se. Indeed, it is reported that PIS is present in a mobile ER-derived subcompartment that makes transient contacts with other organelles, including the plasma membrane, and facilitates the distribution of phosphatidylinositol to other subcellular compartments. The other product of the reaction is cytidine monophosphate (CMP). As PIS catalyzes the reverse reaction also, the rate of phosphatidylinositol synthesis is determined by the relative concentrations of the precursors and product, and the latter must be transported away from the site of synthesis for the reaction to continue. Much of the phosphatidylinositol is delivered to other membranes by vesicular transport, but a family of soluble phosphatidylinositol transfer proteins (PITPα, PITPβ and PITPNC1) provides phosphatidylinositol from the ER to kinases for phosphorylation (see below). Molecular species specificity: The phosphatidylinositol synthase per se does not exhibit the fatty acyl specificity observed in the final product, but earlier in the biosynthetic process 1-stearoyl-2-arachidonoyl species of diacyl-sn-glycerols are converted preferentially into phosphatidic acid by the epsilon isoform of diacylglycerol kinase (DGKε), anchored to the membrane via its N-terminal hydrophobic helix segment; ATP is the phosphate donor. In addition, one of the CDP-diacylglycerol synthases (CDS2) has similar specificity in the generation of the immediate precursor CDP-diacylglycerols from phosphatidic acid, while some specificity may be introduced via lysophosphatidylinositol, formed as a by-product of eicosanoid formation (see below) or as an intermediate as part of the normal cycle of deacylation-acylation of phosphatidylinositol in tissues in which the fatty acid composition is remodeled to give the final distinctive composition. A membrane-bound O-acyltransferase (MBOAT7 or LPIAT1) specific for position sn-2 of lysophosphatidylinositol with a marked preference for arachidonoyl-CoA is ubiquitously expressed in animal tissues, and this may be one means by which free arachidonic acid and eicosanoid levels are regulated. In macrophages subjected to inflammatory stimuli, phosphatidylinositol containing two molecules of arachidonate is produced by remodeling reactions, and there is evidence that it is a novel bioactive phospholipid regulating innate immune responses in these cells. Further specificity may be introduced by lysocardiolipin acyltransferase (LYCAT; also known as LCLAT1 or ALCAT1), which exhibits a preference for lysophosphatidylinositol and lysophosphatidylglycerol over other phospholipids in vitro, and incorporates 18:0 rather than shorter chain fatty acids into position sn-1 of phosphatidylinositol and other phosphoinositides, especially phosphatidylinositol-4,5-bisphosphate and phosphatidylinositol-3-phosphate; this enzyme may be located adjacent to the phosphatidylinositol synthase in the endoplasmic reticulum. Some of the phosphatidylinositol in membranes is derived from recycling of polyphosphoinositides via the phosphatidylinositol cycle, and this could influence the molecular species composition (see below). The highly specific distribution of fatty acids on the glycerol moiety of phosphatidylinositol breaks down in some cancer cells, especially those with a mutation on the transcription factor p53 gene, which is one of the most highly mutated genes in cancers. Plants and bacteria: In contrast to animals, plants have two phosphatidylinositol synthase isoforms, PIS1 and PIS2, which display specificities for particular species of the CDP-diacylglycerol substrate. PIS1 generates phosphatidylinositol with saturated or monounsaturated fatty acids preferentially, while PIS2 generates polyunsaturated species, the two forms possibly having different functions. In protozoan parasites, such as Trypanosoma brucei, the active site of phosphatidylinositol synthase may be the lumen of the endoplasmic reticulum and Golgi. There is evidence for two distinct pools of product in this organism, the bulk membrane form derived from inositol imported from the environment, and a second used for the synthesis of GPI anchors, which uses myo-inositol synthesized de novo. In yeasts, some biosynthesis may occur on the cytosolic side of the plasma membrane. The enzyme is a transmembrane protein.and use CDP-diacylglycerol as a donor and either inositol (eukaryotes) or inositol phosphate (prokaryotes) as the acceptor alcohol. The structure of a similar enzyme, phosphatidylinositol-phosphate synthase from Renibacterium salmoninarum, is shown in Figure \(33\). Figure \(33\): Structure and reaction of phosphatidylinositol-phosphate synthase from Renibacterium salmoninarum. Clarke, O., Tomasek, D., Jorge, C. et al. Structural basis for phosphatidylinositol-phosphate biosynthesis. Nat Commun 6, 8505 (2015). https://doi.org/10.1038/ncomms9505. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ Panel A shows the reaction for PIP synthases which involves the transfer of a diacylglycerol-substituted phosphate group (purple/red) from the CDP-DAG donor to the inositol phosphate acceptor (green), generating PIP and CMP. Panel B shows the structure of the RsPIPS-Δ6N homodimer in ribbon representation viewed from two orthogonal orientations (in the plane of the membrane on the left; towards the cytosol down the dimer axis on the right). One protomer is colored grey, and the helices of the other are depicted in spectral coloring, from blue (JM1) to red (TM6). The Af2299 extramembrane domain used to facilitate crystallization is not shown here. Figure \(34\) shows a large cavity that contains the active site of RsPIPS. Panel (a) shows the structure of RsPIPS-Δ6N is shown in ribbon representation, with one protomer colored grey and the other colored by the Kyte–Doolitle hydrophobicity scale, from −4.5 (most polar, light blue) to 4.5 (most hydrophobic, orange). Two orthogonal representations are shown, on the left is a view in the plane of the membrane, and on the right is a view from the cytosol along the dimer axis. A transparent purple surface delineates the borders of the interfacial cavity, which contains three subregions as follows: 1, the inositol phosphate acceptor-binding pocket; 2, the nucleotide-binding pocket between TM2 and TM3; and 3, a hydrophobic groove between TM2 and JM1. (b) Detail of the active site viewed in the plane of the membrane, with side chains that contact the bound Mg2+ and SO42- ions labeled and depicted in stick representation. A nucleotide-binding site formed from transmembrane segments 1, 2, and 3 contains 8 conserved residues (D1xxD2G1xxAR…G2xxxD3xxxD4). The first 3 aspartic acid side chains coordinate a metal ion while the 4th is likely a general base in catalysis. Figure \(35\) shows an interactive iCn3D model of the phosphatidylinositolphosphate (PIP) synthase with bound CDP-DAG from Renibacterium salmoninarum (5D92). Figure \(35\): Phosphatidylinositolphosphate (PIP) synthase with bound CDP-DAG from Renibacterium salmoninarum (5D92).. Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...i9nhSw9U32Man8 The two identical subunits of the homodimer are shown in gray and plum. The two CDP-DAGs are shown in spacefill and CPK colors Function: In addition to functioning as negatively charged building blocks of membranes, the inositol phospholipids (including the phosphatidylinositol phosphates or 'polyphosphoinositides' discussed below) have crucial roles in the interfacial binding of proteins and in the regulation of protein activity at the cell interface. As phosphoinositides are polyanionic, they can be very effective in non-specific electrostatic interactions with proteins. However, they are especially efficient in specific binding to so-called ‘PH’ domains of cellular proteins. At least three phosphatidylinositol molecules are present in the crystal structure of human erythrocyte glycophorin, for example, and they are believed to influence binding to other proteins via their head groups. The lipid is a structural component of yeast cytochrome bc1. In animal tissues, phosphatidylinositol is the primary source of the arachidonic acid required for the biosynthesis of eicosanoids, including prostaglandins, via the action of the enzyme phospholipase A2, which releases the fatty acids from position sn-2. The reverse reaction also occurs. Figure \(36\)s shows the generation of arachidonic acid and eicosanoids from PI by means of phospholipase A2 Similarly, phosphatidylinositol and the phosphatidylinositol phosphates are the main sources of diacylglycerols that serve as signaling molecules in animal and plant cells, via the action of a family of highly specific enzymes collectively known as phospholipase C. In brief, diacylglycerols regulate the activity of a group of at least a dozen related enzymes known as protein kinase C, which in turn control many key cellular functions, including differentiation, proliferation, metabolism, and apoptosis. Indeed, the biological actions of the various components released have been the subject of intensive study over many years. 2‑Arachidonoylglycerol, an endogenous cannabinoid receptor ligand, may also be a product of phosphatidylinositol catabolism. Phosphatidylinositol Phosphates (Polyphosphoinositides) in Animals Structure and Occurrence: The pioneering work of Mable and Lowell Hokin in the 1950s led to the discovery that phosphatidylinositol was converted to polyphosphoinositides with important signaling and other functional activities, including cell communication via signal transduction, cell survival and proliferation, membrane trafficking and modulation of gene expression. Phosphatidylinositol is now known to be phosphorylated by a number of substrate-selective kinases that place the phosphate moiety on positions 3, 4, and 5 of the inositol ring with the balance among them maintained by distinct phosphatases and phospholipases. Seven different isomers are known (mono-, bis-, and tris-phosphorylated), which are produced in a tightly coordinated manner, and all of these have characteristic biological activities. They each turn over much more rapidly than the parent phosphatidylinositol molecule. In addition, there can be an array of molecular species of each of these isomers that differ in the nature of the fatty acyl groups. Although the most significant in quantitative and possibly biological terms were long thought to be phosphatidylinositol 4-phosphate and phosphatidylinositol 4,5‑bisphosphate, it is now recognized that phosphatidylinositol 3-phosphate and its metabolites are as important biologically at least. Figure \(37\) shows the structures of phosphatidylinositol phosphates These lipids are usually present at low levels only in tissues, typically at about 0.5 to 1% of the total lipids of the inner leaflet of the plasma membrane, so they are unlikely to have an appreciable structural role. On the other hand, static measurements of lipids that turn over very rapidly do not provide a meaningful assessment of their cellular functions. The positional distributions of fatty acids in the phosphatidylinositol, phosphatidylinositol 4-phosphate, and phosphatidylinositol 4,5-bisphosphate of ox brain are listed in Table \(7\)​​​​​​​. In each the saturated fatty acids are concentrated in position sn-1 and polyunsaturated, especially arachidonate, in position sn-2; there are few differences among the three lipids in this instance. Table \(7\)​​​​​​​: Distribution of fatty acids (mol % of the total) in positions sn‑1 and sn‑2 in phosphatidylinositol (PI) and the phosphatidylinositol mono- and diphosphates of ox brain. Fatty acids PI PI monophosphate PI diphosphate sn-1 sn-2 sn-1 sn-2 sn-1 sn-2 16:0 15   9   7 18:0 74   69   69 18:1 10 10 20 13 21 10 18:2 1 2 trace 1 1 1 20:3(n-9)   5   10   10 20:3(n-6)   5   11   12 20:4(n-6)   67   49   52 22:3   7   10   7 22:6(n-3)   trace   trace   trace Data from Holub, B.J. et al., J. Lipid Res., 11, 558-564 (1970); DOI. Molecular species data, see Traynor-Kaplan, A. et al., Biochim. Biophys. Acta, 1862, 513-522 (2017); DOI. Biosynthesis: Phosphatidylinositol per se is the ultimate precursor of all phosphoinositides, the head groups of which have different charges and structures that impact directly on membrane properties and via metabolic interactions can function as chemical switches. The individual phosphoinositides are maintained at steady state levels in membranes by a continuous and sequential series of phosphorylation and dephosphorylation reactions by specific kinases, phosphatases, and phospholipase C enzymes, which are regulated and/or relocated through cell surface receptors for extracellular ligands, the phosphoinositide cycle. While this has been termed a ‘futile cycle’, which can consume a significant proportion of cellular ATP production, it is only part of a wider pattern of reactions - the phosphatidylinositol cycle (see below). Controlled synthesis of these different phosphoinositides occurs in different intracellular compartments for distinct and independently regulated functions with spacially distinct target enzymes or receptors. In mammals, the complexity is such that 18 phosphoinositide inter-conversion reactions have been identified to date, and these are mediated by at least 20 phosphoinositide kinases and 34 phosphoinositide phosphatases that span 8 and 10 classes, respectively; some have yet to be characterized. Most of these enzymes are conserved across all of the eukaryota, and each has distinct functions and specificities that cannot be replaced by the activity of related isoforms. As a generality, most mono-phosphorylations occur in endomembranes, such as the endosomes and the Golgi network, while second and third phosphorylations occur primarily at the plasma membrane, and this is reflected in the lipid composition of each membrane. While these enzymes are believed to work independently and sequentially to produce a specific product, there remains a possibility that some participate in protein complexes to coordinate their activities. Specific transporters, especially the 'Nir2' protein, facilitate the exchange of phosphoinositides between membranes. It should be noted that there are links to the metabolism of phosphatidylcholine, which can be hydrolyzed by phospholipase D to phosphatidic acid, an important activator of key kinases. Figure \(38\) shows an overview of polyphosphoinositide metabolism in animal tissues. Thus as an example, phosphatidylinositol 4-phosphate (PI(4)P) is produced by the action of a phosphatidylinositol 4-kinase (PI4K) in the Golgi, and is in turn phosphorylated by a phosphatidylinositol phosphate 5-kinase (PIPK I) to form phosphatidylinositol 4,5-bisphosphate (PI(4,5)P) at the plasma membrane, although this can also be formed by phosphorylation of phosphatidylinositol 5-phosphate by a specific 4-kinase (PIPK II). Four isoforms of PI4K in two structural families are known that each operate in different subcellular membrane compartments to produce phosphatidylinositol 4-phosphate for particular signaling functions. Some selectivity in the formation of molecular species or remodeling may occur to further enrich the arachidonic acid content. Subsequently, it was discovered that phosphatidylinositol is also phosphorylated by a 3-kinase (PI3K III or the VPS 34 complex) to produce phosphatidylinositol 3-phosphate (PI(3)P) in the early endosomes. In fact, three phosphatidylinositol 3-kinases families (eight isoforms) have been described, each with distinct substrate specificities. A second phosphoinositide signaling pathway involves activation of two of these 3‑kinases, stimulated by growth factors and hormones, which phosphorylate phosphatidylinositol 4,5-bisphosphate (by PI3K I - four isoforms) and phosphatidylinositol 4‑phosphate (by PI3K II - three isoforms) to produce phosphatidylinositol 3,4,5-trisphosphate (PI(3,4,5)P) and phosphatidylinositol 3,4‑bisphosphate (PI(3,4)P), respectively. While phosphatidylinositol 3-phosphate and other 3‑phosphorylated metabolites amount to only about 0.5% of the total phosphoinositides in resting mammalian cells, they are now recognized to be of profound importance for cellular metabolism. In addition to the activity of kinases, the amounts of these various metabolites are regulated by the activities of specific phosphoinositide phosphatases, which are highly conserved in eukaryotes and dephosphorylate phosphoinositides at the 3, 4, and 5 positions of the inositol ring. For example, so-called ‘SHIP’ phosphatases convert phosphatidylinositol 4,5‑bisphosphate back to phosphatidylinositol 4-phosphate by hydrolysis of the 5-phosphate group. 3‑Phosphorylated phosphoinositides are only degraded by phosphatases, especially those of the PTEN family, and not by phospholipase C (see below). The various organelles in cells have membranes with distinct functions and molecular compositions. Yet, all the phosphatidylinositol precursor is formed primarily at the endoplasmic reticulum, and the different membrane lipids must be transported between membrane sites via specific trafficking processes/proteins. There is selective recruitment of effector proteins to particular membranes by binding only to a single type of phosphoinositide, and this is followed by interactions between the phosphoinositide-binding proteins and various enzymes to channel phosphoinositide production to the required biological outcomes and to regulate signaling. For example, much of the phosphatidylinositol 4‑phosphate and phosphatidylinositol 4,5-bisphosphate involved in signaling is believed to be formed at contact sites between the endoplasmic reticulum and plasma membrane. A concept has emerged in which each phosphoinositide has its own role – the ‘lipid code’ hypothesis, in which defined lipids act as labels for each cellular membrane to organize cells into dynamic and responsive membrane-bound compartments and maintain the orderly flow required for the complexities of membrane trafficking and spatio-temporal signaling reactions. Thus, phosphatidylinositol 4-phosphate, phosphatidylinositol 4,5‑bisphosphate, phosphatidylinositol 3-phosphate and phosphatidylinositol 3,5‑bisphosphate are found mainly on the Golgi, plasma membrane, early endosomes, and late endocytic organelles, respectively, where they are sometimes regarded as landmarks for these compartments. For example, phosphatidylinositol 4,5‑bisphosphate is present throughout the plasma membrane and is considered a general marker for this, while phosphatidylinositol 3,4,5-triphosphate, is a characteristic component of the basolateral region of this membrane in a polarized cell but is absent from the apical part. On the other hand, it should be noted that this map of phosphoinositides to specific organelles is derived from their steady state distributions, but the highly dynamic generation and consumption of different phosphoinositides in response to different stimuli in the various sub-cellular compartments in living cells by the action of kinases and phosphatases together with lipase reactions, may lead to the formation of transient pools of distinct molecular forms. There must be a continuous replenishment of the precursors by new synthesis. Function: The distinctive subcellular location of the different phosphoinositide species, together with the rapid and reversible nature of phosphorylation, gives them a central and general position in the fields of cell signaling cascades and intracellular membrane trafficking. The precise locations of particular phosphoinositides are factors that contribute a specific identity to each organelle and sometimes even to each face of an organelle, such as the cis and trans faces of the Golgi apparatus, and this enables directional transport of cellular constituents between organelles or membranes. Phosphoinositides are able to achieve signaling effects directly by binding to specific cytosolic domains of membrane proteins via their polar head groups, thereby triggering downstream signaling cascades, often in conjunction with an acidic phospholipid, such as phosphatidylserine or phosphatidic acid at an adjacent-binding site. The term 'lipidon' has been coined to describe the unique collection of co-located lipids that distinguish biological membrane nano-environments and which provide the context for PI recognition in vivo. In this way, they can regulate the function of innumerable proteins integral to membranes, for example by relocating a protein from one area of the cell to another, e.g., from the cytosol to the inner leaflet of the plasma membrane, or they can attract cytoskeletal and signaling components to the membrane. Amongst the proteins that bind to phosphoinositides in this way are phospholipases, protein kinases, regulators of membrane trafficking, and cytoskeletal, scaffold, and ion channel proteins. Dysregulation of phosphoinositide metabolism and signaling is a factor in a number of diseases, including cancer. Binding usually involves electrostatic interactions with the negative charges of the phosphate groups on the inositol ring with characteristic clusters of basic amino acid residues in proteins to recruit them to intracellular membranes, while often leading to specific folding and thence increased activity of unstructured peptides. At least 70 distinct types of binding sites for phosphoinositides have been identified in proteins. In particular, a binding region termed the pleckstrin homology (PH) domain, consisting of ~100 amino acids, is the most abundant lipid-binding domain with more than 225 examples identified, and this can exhibit great specificity for particular polyphosphoinositides, often binding simultaneously with other proteins. While the interaction is driven by non-specific electrostatic interactions initially, it is followed by specific binding to increase the membrane residence time. The phox homology (PX) domain family with 49 members in humans is unique in that it can recognize all seven phosphoinositide forms, while proteins with a FYVE domain, which is enriched in cysteine and is stabilized by two zinc atoms, bind specifically to phosphatidylinositol 3-phosphate (PI(3)P). The protein kinase C family have C1 or C2 domains which recognize phosphatidylinositol 4,5-bisphosphate and phosphatidylinositol 3,4,5-trisphosphate specifically (and sometimes other lipids). The distinctive phosphoinositide composition of membranes in different organelles adds strength and specificity to the interactions by cooperative binding with other membrane proteins. Phosphatidylinositol 3-phosphate and the other phosphatidylinositol monophosphates are present in cells at low levels only, although their levels do not appear to fluctuate greatly. PI(3)P has been implicated in membrane trafficking through its interactions with certain proteins in endosomes. In particular, it plays a pivotal role in the initiation of autophagy, i.e. the controlled internal degradation and turnover of cellular constituents, while PI(3,5)P2 is important in the autophagosome–lysosome fusion step and in the subsequent acidification of this organelle. After sorting of the lysosomal contents, components of the internalized cargo are recycled to the plasma membrane and PI(3)P is dephosphorylated to phosphatidylinositol by a specific phosphatase, and this is in turn phosphorylated to PI(4)P. Thus the processes of internalization, sorting, and trafficking of membrane proteins depend on the interconversion of phosphoinositide species by coordinated phosphorylation-dephosphorylation reactions. In general, PI(3)P controls cellular processes by recruiting effector proteins through low to moderate affinity interaction with specific PI(3)P binding domains. A protein designated Akt (protein kinase B) is recognized as a direct effector of the PI3K signaling cascade with receptor tyrosine kinases as the main upstream activators, for example, but it is now known that every phosphatidylinositol phosphate has a specific set of effector proteins that are recruited to target membranes or are allosterically regulated by the specific receptors; each function may require a different effector. A further function of PI(3)P is in the regulation of the final stage of cell division (cytokinesis), and the lipid is known to accumulate where cells divide. As the class I PI3K isoforms especially have been implicated in the etiology and maintenance of various diseases and metabolic disorders, including cancer, inflammation, and autoimmunity, drug companies are actively pursuing the development of inhibitors. In particular, they mediate insulin-independent glucose transport and many of the physiological actions of insulin. In relation to lung cancer especially, RAS proteins, which are key signaling switches essential for the control of proliferation, differentiation, and survival of eukaryotic cells, regulate the activity of type I phosphatidylinositol 3-kinase (PI3K); this is essential for tumor initiation and maintenance. Phosphatidylinositol 4-phosphate is the precursor for the 4,5-bisphosphate, but it binds to a protein on the cytoskeleton of the cell and has its own characteristic functions. It is the most widely distributed of the phosphoinositides, and in addition to the Golgi and the plasma membrane, it is present in late endosomes, lysosomes, secretory vesicles, and autophagosomes. As a part of protein-lipid complexes, it is believed to have a role in essential nuclear processes. In yeast, it has a function in the anterograde transport from the trans-Golgi and the retrograde transport from the Golgi to the endoplasmic reticulum; it is also necessary for the formation of secretory vesicles in the Golgi that are targeted to the plasma membrane. Some of that in the plasma membrane is exchanged for phosphatidylserine by the action of specific transport proteins at junctions with the endoplasmic reticulum. In addition, PI(4)P is essential for the structure and function of the late endosomes, where it is required for the recruitment of specific proteins that control cargo exit (following hydrolysis of PI(3)P). Some of these participate in vesicle formation, while others like the oxysterol binding protein (OSBP) are involved in lipid transfer. After initiation of the process by PI(3)P, PI(4)P, PI(4,5)P2 and their binding proteins are modulators of autophagy at most stages of the process. PI(4)P has been called the 'fuel' that drives cholesterol transport, as its hydrolysis provides the energy that enables the establishment of active sterol concentration gradients across membrane-bound compartments with the aid of OSBP, which is a key regulator of cholesterol, oxysterol, and PI(4)P concentrations in membranes. In the plasma membrane, PI(4)P can support the functions of ion channels, and it contributes to the anchoring of proteins with polybasic domains, although it is not utilized for the synthesis of PI(4,5)P2 in this membrane. On the other hand, PI(4)P derived from PI(4,5)P2 in the membrane of primary cilia in the retina is important for vision. PI(4)P has an important influence on the progression of many diseases, especially virus replication, cancer, and various inflammatory diseases, and inhibitors of PI4-kinase are under study for their therapeutic potential. While the biological properties of phosphatidylinositol 5-phosphate have taken longer to unravel, because of the difficulties of separation of this isomer, it is now apparent that it is involved in osmoregulation both in plants and animals. It also has signaling functions, and although it is the least abundant phosphatidylinositol monophosphate, it is involved in signaling at the nucleus and in the cytoplasm, modulating cellular responses to various stresses, hormones and growth factors. In the endosomes, it is a regulator of protein sorting. Although phosphatidylinositol 4,5-bisphosphate (PI(4,5)P2) is found primarily in the inner leaflet of the plasma membrane, where it may define membrane identity in eukaryotic cells, it is also present in endosomes, the endoplasmic reticulum and nucleus. It is an essential precursor of lipid second messengers such as diacylglycerols with vital signaling functions that operate through plasma membrane G-protein coupled receptors, receptor tyrosine kinases, and immune receptors. Because of its large head group and multivalent negative charge, PI(4,5)P2 has been described as an "electrostatic beacon" that interacts in various ways with membrane proteins, other lipids and cellular cations. In consequence and in spite of its relatively low concentration, it is a key regulator of innumerable events at the plasma membrane, including cell adhesion and motility, vesicle endocytosis and exocytosis, and the function of ion channels, especially those for potassium, calcium, and sodium. With ion channels, for example, it appears to be an obligatory factor, increasing their activity by activating key proteins, while its hydrolysis by phospholipase C reduces such activity. PI(4,5)P2 interacts with cationic residues of a large array of proteins in concert with cholesterol to form localized membrane domains that are distinct from the sphingolipid-enriched rafts. Indeed, it has a much higher concentration than other phosphoinositide species in cells, although most of this is in effect sequestered by binding proteins. Also, phosphatidylinositol 4,5-bisphosphate and its diacylglycerol metabolites are important for vesicle formation in membranes. For example, a major pathway in cells for the internalization of cell surface proteins such as transferrin is the clathrin-coated vesicle pathway. PI(4,5)P2 is essential to this process in that it binds to the machinery involved in the membrane, increasing the number of clathrin-coated pits and permitting the internalization of proteins. It has a related function in caveolae, where it is concentrated at the rim. Through its attachment to the apical plasma membrane, phosphatidylinositol 4,5-bisphosphate is intimately involved in the development of the actin cytoskeleton and thereby controls cell shape, motility, and many other processes. In particular, it binds with high specificity to effectors such as vinculin, a membrane-cytoskeletal protein that is involved in the linkage of integrin adhesion molecules to the actin cytoskeleton. Dysregulation of this function has been implicated in the migration and metastasis of tumor cells. In yeasts, it appears that the presence of stearic acid in position sn-1 is essential for this function. In the cell nucleus, this lipid is believed to be involved in maintaining chromatin, the complex combination of DNA, RNA, and protein that makes up chromosomes in a transcriptionally active conformation, as well as being a precursor for further signaling molecules. It has a role in gene transcription, and RNA processing, especially in the modulation of RNA polymerase activity, and in other nuclear processes. Via its binding to specific proteins, the lipid is an essential component of the immune response of animal tissues to toxic bacterial lipopolysaccharides. It is also involved in the pathophysiology of the HIV virus via an interaction with the Tat protein secreted by infected cells. PI(4,5)P2 is the primary precursor of the endocannabinoid 2-arachidonoylglycerol in neurons, and it is also an essential cofactor for phospholipase D and so affects the cellular production of phosphatidic acid with its specific signaling functions. By binding specifically to ceramide kinase, the enzyme responsible for the synthesis of ceramide-1-phosphate, it has an influence on sphingolipid metabolism. Like ceramide-1-phosphate, it binds to and activates the Ca2+-dependent phospholipase A2, which generates the arachidonate for eicosanoid production. One molecule of phosphatidylinositol 4,5‑bisphosphate is bound to each subunit of the protein in the X-ray crystal structures of mammalian GIRK2 potassium channel, where it enables a conformational change that assists the transport function of the protein. Perhaps, the best characterized of the phosphoinositide signaling functions results from the hydrolysis of phosphatidylinositol phosphates by phospholipase C isoforms, in this instance to produce sn-1,2-diacylglycerols and inositol 3,4,5-trisphosphate (see below), which act as second messengers. Only those polyunsaturated diacylglycerol species derived from PI(4,5)P2 are able to bind and activate protein kinase C (α, ε, δ) isoforms both in vitro and in vivo. This lipid is doubly important as it binds strongly to these enzymes via a basic patch distal to a Ca2+ binding site, and this targets them selectively to the plasma membrane. Aberrant expression of phospholipase Cγ2 may be a factor in neurodegenerative diseases. Via the action of PI3 kinase, PI(4,5)P2 is the precursor of PI(3,4,5)P3 with its own distinctive signaling properties. Phosphatidylinositol 3,4-bisphosphate can be produced by two routes and regulates a variety of cellular processes with relevance to health and disease that include B cell activation and autoantibody production, insulin sensitivity, neuronal dynamics, endocytosis, and cell migration. It is known to bind selectively to a number of proteins, and it acts as a secondary messenger by recruiting the protein kinases Akt (protein kinase B) and so may influence the cell cycle, cell survival, angiogenesis, and glucose metabolism. During endocytosis in the endolysosomal system, it is produced from PI(4,5)P2 and controls the maturation of endocytic-coated pits. Its synthesis and turnover of are spatially segregated within the endocytic pathway. In epithelial cells, it is located on the apical membrane, i.e. facing the lumen, as opposed to the basolateral membranes, and it is believed to be is a determinant of the identity and function of the apical membrane. Phosphatidylinositol 3,5-bisphosphate is present at low levels only in cells (0.04-0.1% of the total phosphatidylinositides) unless stimulated by growth factors, but it is important in membrane and protein trafficking, especially in the late endosomes in eukaryotes and in yeast vacuoles. For example, the conversion of PI(3)P to PI(3,5)P2 promotes endosomal maturation and degradative sorting. It is involved in the mediation of signaling in response to stress and hormonal cues and in the control of ion transport in membranes, while genetic studies confirm that it is essential for healthy embryonic development, especially in the nervous system. Phosphatidylinositol 3,4,5-trisphosphate (PI(3,4,5)P3) is almost undetectable in quiescent cells, but its intracellular level rises very rapidly from synthesis at the plasma membrane in response to agonists such as extracellular growth factors and hormonal stimuli. By recruiting proteins with pleckstrin homology (PH) domains to the plasma membrane, it has been implicated in a variety of cellular functions that include growth, cell survival, proliferation, cytoskeletal rearrangement, intracellular vesicle trafficking, and cell metabolism. In particular, it is an important component of a signaling pathway in the cell nucleus. In epithelial cells, it is located on the basolateral membrane, i.e. facing adjacent cells, where it may be a determinant of the identity and function of this membrane. In contrast to phosphatidylinositol 3-phosphate, it opposes autophagy by binding to and activating the PH domain of Akt, so inducing cell proliferation. During feeding, various physiological responses lead to the secretion of insulin, which stimulates the phosphorylation of phosphatidylinositol 4,5-bisphosphate to phosphatidylinositol 3,4,5-trisphosphate and triggers a signaling cascade that leads to the suppression of autophagy. When this pathway is impaired it has deleterious effects on insulin resistance associated with various metabolic diseases including obesity and diabetes. It has been implicated in tumor cell migration and metastasis. PI(3,4,5)P3 is also present in the nucleus and nucleoli of cells where it is believed to have functions in RNA processing/splicing, cytokinesis, protein folding, and DNA repair. In complete contrast, like phosphatidylserine, it is reportedly transferred to the outer leaflet of the plasma membrane in aged or damaged cells as an 'eat‑me' signal for phagocytes and apoptosis. The human immune system utilizes neutrophils, which are highly mobile cells, to eliminate pathogens from infected tissue. The first step is to track and then pursue molecular signals, such as cytokines, emitted by pathogens. It has been established that two phospholipids operate in sequence to point the neutrophils in the correct direction. The first of these is phosphatidylinositol 3,4,5-trisphosphate, which binds to a specific protein DOCK2 and enables it to translocate to the plasma membrane. Then phosphatidic acid, generated by the action of phospholipase D on phosphatidylcholine, takes over and directs the DOCK2 to the leading edge of the plasma membrane. This causes the polymerization of actin within the cell and in effect reshapes the neutrophil and points it in the direction from which the pathogens signals are coming. On the other hand, Mycobacterium tuberculosis is able to subvert phosphoinositide signaling to arrest phagosome maturation by dephosphorylation of phosphatidylinositol 3-phosphate. Water-Soluble Inositol Phosphates As mentioned briefly above, hydrolysis of phosphatidylinositol phosphates by calcium-dependent phospholipase C (or 'phosphoinositidase C') leads to the generation of sn‑1,2‑diacylglycerols, which act as second messengers in animal cells and are of enormous metabolic importance. There are many different enzymes of this type, but the activity of the phosphoinositide-specific phospholipase C constitutes an essential step in the inositide signaling pathways. The enzyme exists in six families consisting of at least 13 isoenzymes, all of which have conserved regions such as the plekstrin homology (PH) binding domain. Each one has a distinctive role and can have a characteristic cell distribution that is linked to a specific function. The activity of these enzymes is stimulated by signaling molecules such as G-protein coupled receptors, receptor tyrosine kinases, Ras-like GTPases, and calcium ions, thus linking the hydrolysis of phosphatidylinositol phosphates to a wide range of other cellular signals. As phospholipase C is a soluble protein located mainly in the cytosol, translocation to the plasma membrane is a crucial step in signal transduction. Regulation of these isoenzymes and the form PLCγ1 in particular is vital for health as they are associated with the activation or inhibition of important pathophysiological processes, especially in relation to cancer. Some phosphatidic acid is synthesized from the diacylglycerols produced within the plasma membrane through the activity of diacylglycerol kinases, and this is transported back to the endoplasmic reticulum and ultimately can be re-utilized for phosphatidylinositol biosynthesis. The other products of the phospholipase C reaction that are of special relevance because of their many essential functions are water-soluble inositol phosphates. Up to 60 different compounds of this type are possible, and at least 37 of these have been found in nature at the last count, all of which are extremely important biologically. However, polyphosphoinositides with a phosphate in position 3 are not substrates for phospholipase C. Figure \(39\) shows the generation of inositol phosphates by phospholipase C. For example, under the action of various physiological stimuli in animals, including sphingosine-1-phosphate, and acting via various G-protein-coupled receptors, phosphatidylinositol 4,5-bisphosphate in the plasma membrane is hydrolyzed to release inositol 1,4,5-trisphosphate, an important cellular messenger that diffuses into the cytosol and stimulates calcium release from an ATP-loaded store in the endoplasmic reticulum via ligand-gated calcium channels (the diacylglycerols remain in the membrane to recruit and activate members of the protein kinase C family). The increase in calcium concentration, together with the altered phosphorylation status, activates or de-activates many different protein targets, enabling cells to respond in an appropriate manner to the extracellular stimulus. To enable rapid replenishment of the phosphatidylinositol 4,5‑bisphosphate used in this way, a cycle of reactions - the phosphatidylinositol cycle - must occur (see below). On the other hand, a recent publication suggests that phosphatidylinositol 4-phosphate in the plasma membrane may be a more important source of diacylglycerols following stimulation of G protein–coupled receptors. All of the various inositol phosphates appear to be involved in the control of cellular events in very specific ways, but especially in the organization of key signaling pathways, the rearrangement of the actin cytoskeleton, or intracellular vesicle trafficking. They have been implicated in gene transcription, RNA editing, nuclear export, and protein phosphorylation. As these remarkable compounds can be rapidly synthesized and degraded in discrete membrane domains or even sub-nuclear structures, they are considered to be ideal regulators of dynamic cellular mechanisms. From structural studies of inositol polyphosphate-binding proteins, it is believed that the inositides may act in part at least by modifying protein function by acting as structural cofactors, ensuring that proteins adopt their optimum conformations. In addition, phosphoinositides and the inositol polyphosphates are key components of the nucleus of the cell, where they have many essential functions, including DNA repair, transcription regulation, and RNA dynamics. It is believed that they may be activity switches for the nuclear complexes responsible for such processes, with the phosphorylation state of the inositol ring being of primary importance. As different isomers appear to have specific functions at each level of gene expression, extracellular events must coordinate the production of these compounds in a highly synchronous manner. In organisms from plants to mammals, an extra tier of regulatory mechanisms is produced by kinases that generate energetic diphosphate (pyrophosphate)-containing molecules from inositol phosphates. Conversely, these can be dephosphorylated by polyphosphate phosphohydrolase enzymes to regenerate the original inositol phosphates. These inositol pyrophosphates and the enzymes involved in their metabolism are also involved in the regulation of cellular processes by modulating the activity of proteins by a variety of mechanisms. It should be noted that the phospholipase C isoenzymes regulate the concentration of phosphatidylinositol 4,5-bisphosphate and related lipids and thence their activities in addition to the generation of new biologically active metabolites. Phosphatidylinositides in Plants In plants as in animals, phosphatidylinositol and polyphosphoinositides have essential biological functions, exerting their regulatory effects by acting as ligands that bind to protein targets via specific lipid-binding domains and so alter the location of proteins and their enzymatic activities. However, it appears that polyphosphoinositide metabolism developed in different ways after the divergence of the animal and plant kingdoms so the details of the processes in each are very different, not least because the subcellular locations of phosphoinositides differ appreciably between plants and animals. Phosphatidylinositol per se is of course the precursor of the phosphorylated forms and determines their fatty acid compositions. It also has a role in inhibiting programmed cell death by acting as the biosynthetic precursor of the sphingolipid ceramide phosphoinositol and so reducing the levels of ceramide. As in animals, the various phosphoinositides (five in total) are produced and inter-converted rapidly by a series of kinases and phosphatases (in many isoforms) in different cellular membranes in response to environmental or developmental cues. For example, phosphatidylinositol is generated mainly in the endoplasmic reticulum, while PI 4-kinases and their product are located in the trans-Golgi network and nucleus, and PI4P 5-kinases and product are present in the plasma membrane. During the biosynthesis of polyphosphoinositides, the first phosphorylation occurs at the hydroxyl group at positions 3 or 4 of the inositol ring, catalyzed by the appropriate kinases, while the second phosphorylation then takes place at position 5; PI 5-phosphate is produced by the action of a phosphatase on PI 3,5‑bisphosphate. Most other metabolites are produced via phosphatidylinositol 3-phosphate, and reports that some phosphatidylinositol 3,4,5-trisphosphate may be produced from phosphatidylinositol 4,5‑bisphosphate require confirmation. In contrast to mammalian phosphatidylinositol 3-kinases, which accept both phosphatidylinositol and its monophosphates as substrates, the plant enzyme acts only on the former. Figure \(40\) shows polyphosphoinositide metabolism in plants The reverse reaction in plants is accomplished by phosphoinositide phosphatases, which can be grouped into three main families, the phosphatase/tensin (PTEN) family, 5-phosphatases (5-PTases) and phosphatases containing Suppressor of Actin (SAC) domains, each with differing subcellular locations, substrate specificities and regulatory mechanisms. Although what might be considered normal levels of phosphatidylinositol 4-phosphate are present, the concentrations of phosphatidylinositol 4,5‑bisphosphate and other phosphoinositides are extremely low in plants (10 to 20-fold lower than in mammalian cells), although they still have vital functions. There are differences between cell types, but in Arabidopsis epidermal root cells, PI(4,5)P2 is present at the highest concentration in the plasma membrane (apex region) and nucleus, while PI4P slowly distributes between the plasma membrane and Golgi, with the highest concentration in the former. Multivesicular bodies/late endosomes accumulate both PI3P and PI(3,5)P2, and the tonoplast and autophagosomes contain PI3P. How the various metabolites are transported between membranes has yet to be determined, but non-vesicular transport is believed to occur at membrane contact sites and vesicular transport probably occurs also. Highly polarized distributions of phosphoinositides are found within membranes, generally oriented toward the cytosolic leaflet, and they are believed to be organized in nanoclusters together with other lipids and proteins. For example, phosphatidylinositol-4-phosphate is an important constituent of the plasma membrane in plant cells, where it controls the electrostatic state and is involved in cell division. It may control the location and function of many membrane proteins, including those required for development, reproduction, immunity, nutrition, and signaling. PI(4)P is the only phosphoinositide present at the cell plate, i.e. the membrane separating two daughter cells during cell division. In addition, PI(4)P may interact with salicylic acid in the plant immune response, and it is produced during salt stress. However, specific functions are now being discovered for each of the plant phosphoinositides, which are produced rapidly in response to osmotic and heat stress, and it has become evident that a continuous turnover is essential for cell growth and development. For example, they have marked effects on the growth of many cell types and on guard cell function. In the nucleus, proteins have been identified that bind to phosphoinositides via the acyl chains, leaving the head group exposed for enzymatic modifications and signal transduction. Phosphoinositides are of special importance in microdomains at the tip of growing tissues such as the shoot apical meristem, pollen tubes, and root hairs where phosphatidylinositol 4,5-bisphosphate functions in stem cell maintenance and organogenesis. In the plasma membrane, it is enriched in the detergent-resistant component commonly equated with 'rafts'. Although its concentration is low, PI(4,5)P2 has been shown to have signaling functions by binding to a number of different target proteins, which have characteristic binding domains. For example, together with phosphatidic acid, PI(4,5)P2 regulates the activity of a number of actin-binding proteins, which in turn control the activity of the actin cytoskeleton. This has a key role in plant growth, the movement of subcellular organelles, cell division and differentiation, and plant defense. In addition, this lipid exerts control over ion channels, ATPases, and phospholipase C-mediated lipid degradation and the production of further second messengers. It is an important factor in both clathrin-mediated endocytosis and exocytosis. The specificity of the interactions may be dependent on the fatty acid composition of the lipid and on the activity of phosphatidylinositol 4-phosphate 5-kinase. As in animals, phosphoinositides have a role in endosomal sorting but through the central vacuole, which is a plant-specific organelle with both lytic and storage functions. Phosphatidylinositol 3,5-bisphosphate is the least abundant of the phosphoinositides, but it is a crucial lipid for membrane trafficking systems. The PI to PI(3)P to PI(3,5)P2 cascade, the second step requiring a kinase designated FAB1, is required for endosomal sorting events leading to membrane protein degradation or retrieval, vacuolar morphogenesis and autophagy. PI(3,5)P2 is involved in stomatal closure and the growth of root hairs, and it is also induced in salt stress. A number of different enzymes of the phospholipase C type that are specific for polyphosphoinositides have been isolated from higher plants; they are activated by Ca2+ and unlike their mammalian counterparts, they are not regulated by G proteins. It is not certain whether phosphatidylinositol is itself a substrate for these enzymes in vivo. Less is known of the metabolism of the water-soluble inositol phosphates produced in comparison to animals, and plants appear to lack a receptor for inositol 1,4,5-trisphosphate (IP3), although it is the most abundant metabolite of this type and is reported to induce the release of calcium ions to trigger stomatal closure. However, there is increasing evidence for lipid signaling mediated by phospholipase C in abiotic stress tolerance and development in plants. There is a general if contested belief that inositol hexakisphosphate (phytic acid or IP6), produced at least in part by sequential phosphorylation of inositol 1,4,5-trisphosphate, is a more important cellular messenger in plants and mobilizes an endomembrane store of calcium ions. Inositol-1,2,4,5,6-pentakisphosphate (IP5) is a structural co-factor of the jasmonic acid receptor coronatine insensitive 1, linking phosphoinositide signaling with phytohormone-controlled pathways. In plants in contrast to animals, diacylglycerols, the other product of phospholipase C hydrolysis of phosphoinositides, are rapidly converted to phosphatidic acid by diacylglycerol kinases and have not been considered important in signal transduction. Plants lack protein kinase C but they do have proteins with related properties that appear to be influenced by diacylglycerols. Via the action of phospholipase D, inositol phospholipids are a source of phosphatidic acid with its well-characterized signaling functions in plants, especially in defence. Lyso-Phosphoinositides Figure \(41\) shows the structure of lysophosphatidylinositol Lysophosphatidylinositols: Lysophosphatidylinositols (LPI), i.e. with a single fatty acid only linked to the glycerol moiety, are formed as intermediates in the remodeling of the fatty acid compositions of the lipids by the action of phospholipase A1 or phospholipase A2 (e.g. cPLA2α), and when arachidonic acid is released for eicosanoid biosynthesis (see above). In ovarian cancer, LPI is elevated appreciably to around 15µM in ascites, and it is also present at high levels in obese subjects. It has become apparent relatively recently that like other lysophospholipids, phosphatidylinositol, and polyphospho-analogues may have messenger functions. For example, it has long been known to stimulate the release of insulin from pancreatic cells, suggesting a role in glucose homeostasis. sn-2-Arachidonoyl-lysophosphatidylinositol, in particular, is an endogenous ligand for a G protein-coupled receptor GPR55, and thereby can induce rapid phosphorylation of certain enzymes, including a protein kinase, which promote cancer cell proliferation, migration, and metastasis. Indeed, lysophosphatidylinositol is a biomarker for poor prognosis in cancer patients, and its concentration is elevated significantly in highly proliferative cancer cells in vitro. GPR55 is expressed in many regions of the brain, the intestines, endocrine pancreas and islets (where it may stimulate insulin release). It has been implicated in macrophage activation and inflammation. In addition to its role in cancer, lysophosphatidylinositol has been implicated in a number of metabolic diseases. It is reported to be a precursor of the endocannabinoid 2‑arachidonoylglycerol by the action of human glycerophosphodiesterase 3 as a lysophospholipase C. This enzyme suppresses the receptor for lysophosphatidylinositol, and so acts as a switch between GPR55 and endocannabinoid (CB2) signaling. Glycerophosphoinositol: Sequential removal of both fatty acids from phosphatidylinositol by a specific phospholipase A2 (PLA2IVα) with both phospholipase A2 and lysophospholipase activities releases water-soluble glycerophosphoinositol. While this can be hydrolyzed by a glycerophosphodiester phosphodiesterase to inositol 1-phosphate, glycerophosphoinositol per se has distinctive biological activities and functions, as do related compounds derived from the phosphatidylinositol phosphates. In particular, glycerophosphoinositol has anti-inflammatory activity in that it inhibits the inflammatory and thrombotic responses induced by bacterial lipopolysaccharides (endotoxins). Figure \(42\) shows the structure of glycerophosphoinositol The Phosphatidylinositol Cycle Phosphatidylinositol can be considered to be at the center of a cycle of reactions and intermediates that are involved in innumerable aspects of cellular signaling in animals (a similar cycle could be described for plants). These are discussed individually at length above, but it is useful to point out how each component forms part of a larger pattern. In brief as illustrated, the various synthetic and hydrolytic reactions involved in phosphoinositide metabolism can be considered to constitute a phosphatidylinositol cycle with enzymes located both in the endoplasmic reticulum and plasma membrane, so lipids have to be transferred across the cytosol in both directions between the two to complete the cycle, probably via adjacent membrane structures and facilitated by proteins of the phosphatidylinositol transfer protein membrane-associated family (PITPNM or nir2), which may channel phosphoinositide production to specific biological outcomes. Phospholipase C and phosphatidylinositol-4-phosphate 5-kinase (PI4P 5K) are located in the plasma membrane, while the cytidine diphosphate-diacylglycerol synthase (CDS2) and phosphatidylinositol synthase are in the endoplasmic reticulum. The epsilon isoform of diacylglycerol kinase (DGKε) is located at contact sites between the endoplasmic reticulum and plasma membrane, but there are nine further isoforms with differing cellular and subcellular locations that may be involved in the cycle. Each turn of the cycle uses a great deal of energy and consumes three moles of ATP, together with cytidine triphosphate and inositol. If it is assumed that the pyrophosphate is hydrolyzed by endogenous pyrophosphatases to inorganic phosphate, the cycle can proceed in one direction only. Figure \(43\) shows the phosphatidylinositol cycle. Factors such as membrane curvature must be taken into account, and the diagram is of necessity a considerable over-simplification. In addition to participating in this cycle, many of the lipid intermediates can be precursors for other lipids, and for example, diacylglycerols are potential precursors for triacylglycerols, while phosphatidic acid is a precursor for phosphatidylcholine and phosphatidylethanolamine. Each lipid intermediate is subject to remodeling of the acyl chains via the Lands cycle, and polyunsaturated fatty acids released can be utilized for eicosanoid production. A further by-product of the cycle is inositol triphosphate, which contributes to the regulation of intracellular calcium levels. It has been suggested that the unique molecular species composition of phosphoinositides (18:0-20:4) could influence their selective recycling back into phosphatidylinositol as many of the enzymes involved have a preference for this substrate. A further proposal is that the phosphatidylinositol cycle could act to enrich this species through multiple passages around the cycle.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/21%3A_Lipid_Biosynthesis/21.03%3A_Biosynthesis_of_Membrane_Glycerolipids.txt

Search Fundamentals of Biochemistry by William (Bill) W. Christie and Henry Jakubowski. This section is an abbreviated and modified version of material from the Lipid Web, an introduction to the chemistry and biochemistry of individual lipid classes, written by William Christie. Sphingolipids Introduction: The sphingolipids comprise a wide range of complex lipids in which the defining component is a long-chain or sphingoid base, which in living tissues is usually linked to a fatty acid via an amide bond. J.L.W. Thudichum, a German chemist working in London, first coined the root term “sphingo-” in 1884 following his discovery of the first glycosphingolipids, because the enigmatic nature of the molecules reminded him of the riddle of the sphinx. Regretfully, the importance of his work was not recognized until 25 years after his death, and it was 1947 before the term “sphingolipide” was introduced by Herbert Carter and colleagues. While they are much less enigmatic than they once were, sphingolipids are extremely versatile molecules that continue to fascinate as new knowledge is gained of their functions in healthy (and diseased) animal and plant tissues. They are found in only a few bacterial genera, but they are present in Sphingomonas, Sphingobacterium and a few other species, and many pathogenic species utilize host sphingolipids to promote infections. Novel sphingolipid structures continue to be reported, and as an example at the last count, 188 of the complex sphingolipids classified as gangliosides, with variations in the complex carbohydrate component alone, had been characterized in vertebrates. Long-chain or sphingoid bases, of which sphingosine is typical, are the basic elements and are the simplest possible functional sphingolipids. They vary in chain length and in the presence of various functional groups including double bonds of both the cis- and trans-configuration at different locations in the aliphatic chain. Ceramides, which contain sphingoid bases linked to fatty acids by amide bonds, vary appreciably in the compositions of both aliphatic components, depending on their biological origins. The structure of sphingosine and ceramide, the sphingolipid building blocks, are shown in Figure \(1\). Long-chain bases and ceramides have important biological properties in their own right, for example in relation to intra- and inter-cellular molecular signaling, especially in animal cells, while another relatively simple sphingolipid, sphingosine-1-phosphate, is now recognized as a key factor in countless aspects of animal metabolism. The concentrations of these bioactive lipids respond rapidly to the action of specific stimuli and then regulate downstream effectors and targets. Ceramides are the precursors of a multitude of sphingo-phospho- and sphingo-glycolipids with an immense range of functions in tissues. The properties and functions of these complex sphingolipids are quite distinct from those of the comparable glycerophospho- and glyceroglycolipids. For example in animals, sphingomyelin has structural similarities to phosphatidylcholine, but has very different physical and biological properties, while the complex oligoglycosylceramides and gangliosides(glycosphingolipids, of which glucosylceramide is the precursor, have no true parallels among the glyceroglycolipids. Figure \(2\) shows the structure of the complex sphingolipids sphingomyelin and glucosylceramide. Complex sphingolipids are synthesized in the endoplasmic reticulum and Golgi, but are located mainly in the plasma membrane of most mammalian cells where they have a structural function and also serve as adhesion sites for proteins from the extracellular tissue. The glycosphingolipids are especially important for myelin formation in the brain. However, sphingolipids have intracellular functions in all cellular compartments, including the nucleus. The first five carbon atoms of the sphingoid base in sphingolipids have a highly specific stereochemistry and constitute a key feature that has been termed the ‘sphingoid motif’, which in comparison to other lipid species facilitates a relatively large number of noncovalent interactions with other membrane lipids, via hydrogen-bonding, ion-ion interactions and induced dipole-induced dipole interactions. A distinctive property of sphingolipids in membranes is that they spontaneously form transient nanodomains termed 'rafts', usually in conjunction with cholesterol, where such proteins as enzymes and receptors congregate to carry out their signaling and other functions. Thus, in addition to their direct effects on metabolism, sphingolipids affect innumerable aspects of biochemistry indirectly via their physical properties. While it may be obvious that a well-balanced sphingolipid metabolism is important for health in animals, increasing evidence has been acquired to demonstrate that impaired sphingolipid metabolism and function are involved in the pathophysiology of many of the more common human diseases. These include diabetes, various cancers, microbial infections, Alzheimer's disease and other neurological syndromes, and diseases of the cardiovascular and respiratory systems. In humans, a number of important genetic defects in sphingolipid metabolism or sphingolipidoses have been detected, especially storage diseases associated with the lysosomal compartment where sphingolipids are catabolized. Sphingolipids and their metabolism are therefore likely to prove of ever increasing interest to scientists. There are appreciable differences in sphingolipid compositions and metabolism between animal and plant cells, both with respect to the aliphatic components and especially the polar head groups, although there are also some important similarities. While sphingomyelin is the most abundant sphingolipid in animals, it does not occur in plants and fungi. Although less is known of the role they play in plants, it has become apparent that complex sphingolipids are much more abundant in plant membranes than was once believed, and it is now recognized that they are key components of the plasma membrane and endomembrane system. Some General Comments on Sphingolipid Metabolism The biosynthesis and catabolism of sphingolipids involves a large number of intermediate metabolites, all of which have distinctive biological activities of their own. In animals, the relationships between these metabolites have been rationalized in terms of a ‘sphingomyelin, sphingolipid or ceramide cycle’, as shown in Figure \(3\). Many different enzymes (and their isoforms) are involved, and their activities depend on a number of factors, including intracellular locations and mechanisms of activation. Each of the various compounds in these pathways has characteristic metabolic properties. Thus, free sphingosine and other long-chain bases, which are the primary precursors of ceramides and thence of all the complex sphingolipids, function as mediators of many cellular events, for example by inhibiting the important enzyme protein kinase C. Ceramides are involved in cellular signaling, and especially in the regulation of apoptosis, and cell differentiation, transformation and proliferation, and most stress conditions. In contrast, sphingosine-1-phosphate and ceramide-1-phosphate promote cellular division (mitosis) as opposed to apoptosis, so that the balance between these lipids and ceramide and/or sphingosine levels in cells is critical and necessitates exquisite control in each cellular compartment. Similarly, the ‘structural’ sphingolipids, such as sphingomyelin, monoglycosylceramides, oligoglycosylceramides and gangliosides, all have unique and characteristic biological functions, some of which are due to their physical properties and location within rafts, nanodomains of membranes. Most of the reactions in the sphingomyelin cycle are reversible and the relevant enzymes are located in the endoplasmic reticulum, Golgi, plasma membrane, and mitochondria, but the more complex sphingolipids are catabolized in the lysosomal compartment. Sphingolipids are especially important in providing the permeability barrier in the skin, where they are characterized by the presence of ultra-­long fatty acyl components as well as fatty acyl groups linked to a hydroxyl group at the terminal end of the N‑linked fatty acids (thereby generating a three‑chain rather than a two‑chain molecule). Metabolic pathways that are comparable to those of the sphingomyelin cycle are believed to occur in plants, as shown in Figure \(4\), although they have not been studied as extensively as those in animals. However, sphingolipid metabolites such as sphingosine-1-phosphate (or analogues) have been linked to programmed cell death, signal transduction, membrane stability, host-pathogen interactions and stress responses, for example. Plants also have a unique range of complex sphingolipids in their membranes, such as ceramide phosphorylinositol and the phytoglycosphingolipids, and these are now known to constitute a higher proportion of the total lipids than had hitherto been supposed, although their functions have hardly been explored. While sphingolipids are produced by relatively few bacterial species, sulfono-analogues of long-chain bases and ceramides (capnoids) are produced by some specie. Fatty acid Components of Sphingolipids The fatty acids of sphingolipids are very different from those of glycerolipids, consisting of very-long-chain (up to C26) odd- and even-numbered saturated or monoenoic and related 2(R)-hydroxy components, while even longer fatty acids (C28 to C36) occur in spermatozoa and the epidermis. The dienoic acid 15,18‑tetracosadienoate (24:2(n‑6)), derived from elongation of linoleic acid, is found in the ceramides and other sphingolipids of a number of different tissues, but at relatively low levels. Polyunsaturated fatty acids are only rarely present, although sphingomyelins of testes and spermatozoa are exceptions in that they contain such fatty acids, which are even longer in chain-length (up to 34 carbon atoms) and include 28:4(n‑6) and 30:5(n‑6). Skin ceramides also contain unusual very-long-chain fatty acids, while yeast sphingolipids are distinctive in containing mainly C26 fatty acids. In plants and yeasts, a similar range of chain-lengths occur as in animals, but 2-hydroxy acids predominate sometimes accompanied by small amounts of 2,3‑dihydroxy acids; saturated fatty acids are most abundant, but monoenes are present in higher proportions in the Brassica family (including Arabidopsis) and a few other species. Some fungal species contain monoenoic fatty acids with a trans-3 double bond and/or a hydroxyl group. Figure \(5\) shows typical sphingolipid fatty acids. Very-long-chain saturated and monoenoic fatty acids for sphingolipid biosynthesis are produced from medium-chain precursors by elongases (ELOVL) in the endoplasmic reticulum of cells in mammals, and there is increasing evidence that specific isoforms are involved in the biosynthesis of certain ceramides. For example, ELOVL1 has been linked to the production of ceramides with C24 fatty acids (saturated and unsaturated), while ELOVL4 is responsible for the ultra-long-chain fatty acids in skin. Yeasts possess three elongation enzymes: Elo1 (for medium to long-chain fatty acids), Elo2 (up to C22) and Elo3 (up to C26). The hydroxyl group is believed to add to the hydrogen-bonding capacity of the sphingolipids, and it helps to stabilize membrane structures and strengthen the interactions with membrane proteins. Hydroxylation is effected by a fatty acid 2-hydroxylase in mammals, i.e. an NAD(P)H-dependent monooxygenase, which is an integral membrane protein of the endoplasmic reticulum. It converts unesterified long-chain fatty acids to 2‑hydroxy acids in vitro and probably also in vivo. For example, experimental evidence has been obtained that is consistent with 2‑hydroxylation occurring at the fatty acid level prior to incorporation into ceramides in the brain of mice where the enzyme is expressed at high levels. A second enzyme of this kind is known to exist but has yet to be characterized, and it is possible that a proportion of the odd-chain fatty acids in brain are synthesized by Peroxisomal α-oxidation of the 2‑hydroxy acids. Similarly, in skin, 2‑hydroxy and non-hydroxy fatty acids as their CoA esters are used with equal facility for ceramide biosynthesis by ceramide synthases. As mutations in the fatty acid 2‑hydroxylase in humans and mice give rise to demyelination disorders, such as leukodystrophy, it is evident that sphingolipids containing 2‑hydroxy acids have unique functions in membranes that cannot be substituted by non-hydroxy analogues. In plants, it appears that 2‑hydroxyl groups are inserted into fatty acyl chains while they are linked to ceramide, as ceramide synthase does not accept hydroxy fatty acids in vitro at least. Two fatty acid 2‑hydroxylases (di-iron-oxo enzymes) have been found in Arabidopsis, with one specific for very-long-chain fatty acids and one for palmitic acid. In fungi, a hydroxyl group is inserted at C2 of the fatty acid in a dihydroceramide intermediate. Although the fatty acids are only occasionally considered in terms of the biological functions of sphingolipids, their influence is considerable, especially but not only in relation to their physical properties and function in membranes. For example, very-long-chain fatty acids may play a role in stabilizing highly curved membrane domains as is required during cell division. The hydrophobic nature of the fatty acyl groups (together with the long-chain bases) enables the hydrogen bonding that is essential for the formation of raft nanodomains in membranes. As a general rule, lipid bilayers containing sphingolipids with 2-hydroxy-fatty acyl or 4-hydroxy-sphingoid base moieties, tend to generate condensed and more stable gel phases with higher melting temperatures than their non-hydroxylated equivalents, because they have a more extended and strengthened intermolecular hydrogen bonding network. Changes in fatty acid composition are seen in some disease states, and for example increased concentrations of fatty acids >C24 are a feature of adrenoleukodystrophy, an X-linked genetic disorder. Removal of very-long-chain fatty acids from sphingolipids in mutants of the model plant Arabidopsis inhibits completely the development of seedlings. As example of a more specific interaction, it has been demonstrated that synthetic glycerolipids must contain very-long-chain fatty acids (C26) to allow growth in yeast mutants lacking sphingolipids, probably by stabilizing the proton-pumping enzyme H+-ATPase. Similarly, ceramides containing different fatty acids can be used in highly specific ways. Thus in fungi, C16 or C18 hydroxy acids are used exclusively for synthesis of glucosylceramide, while those containing very-long-chain C24 and C26 hydroxy acids are used only for synthesis of glycosyl inositol phosphorylceramide anchors for proteins. In plants, sphingolipids containing 2-hydroxy acids are protective against oxidative and other biotic stresses. Links between Glycerolipid and Sphingolipid Metabolism Sphingolipid metabolism and glycerolipid metabolism have been widely treated as separate sciences until relatively recently, partly for historical reasons and partly because the analysis of the two lipid groups required different approaches and skills. However, there are many areas where the two overlap, not least because phosphatidylcholine is the biosynthetic precursor of sphingomyelin in animal cells, while in plants and fungi, phosphatidylinositol is the biosynthetic precursor of ceramide phosphorylinositol. In contrast, ethanolamine phosphate derived from the catabolism of sphingolipids via sphingosine 1-phosphate is recycled for the biosynthesis of phosphatidylethanolamine, and this is essential for survival in the protozoan parasite Trypanosoma brucei. In studies in vitro, sphingosine 1-phosphate has been shown to be an activator of the phospholipase C involved in the hydrolysis of the lipid mediator phosphatidylinositol 4,5-bisphosphate with formation of diacylglycerols and inositol triphosphate. The location and functions of glycerophospholipids in membranes is influenced both positively and negatively by sphingolipid-rich domains or rafts in membranes. In addition, there are several examples of phosphoinositides and other complex lipids binding to enzymes of sphingolipid metabolism, either as part of a regulatory function that controls their activity or to facilitate their location to various membranes. Thus, sphingosine kinase 2, one of the enzymes responsible for the biosynthesis of sphingosine 1-phosphate, binds to phosphatidylinositol monophosphates, while the ceramide kinase responsible for the biosynthesis of ceramide 1-phosphate requires phosphatidylinositol 4,5-bisphosphate to function. Similarly, the CERT protein involved in ceramide transport has a binding site for phosphatidylinositol 4-phosphate. Sphingomyelin production at the trans-Golgi network triggers a signaling pathway leading to dephosphorylation of phosphatidylinositol 4-phosphate, interrupting transport of cholesterol and sphingomyelin. Again, the interactions are not solely in one direction as ceramide 1‑phosphate (with phosphatidylinositol 4,5-bisphosphate) binds to the specific phospholipase A2 (cPLA2α) responsible for the hydrolysis of phosphatidylinositol and thence the release arachidonic acid for eicosanoid production. Other than the phosphoinositides, phosphatidylserine activates the neutral sphingomyelinase in brain. Long-Chain (Sphingoid) Bases Long-chain/sphingoid bases are the characteristic and defining structural unit of the sphingolipids, which are important structural and signaling lipids of animals and plants and of a few bacterial species. These are long-chain aliphatic amines, containing two or three hydroxyl groups, and often a distinctive trans-double bond in position 4. To be more precise, they are 2-amino-1,3-dihydroxy-alkanes or alkenes with (2S,3R)‑erythro stereochemistry, often with various further structural modifications in the alkyl chain. They are important for the physical and biological properties of all of the more complex sphingolipids, but free sphingoid bases are also bioactive and interact with specific receptors and target molecules. As discussed below, the mechanisms for biosynthesis of sphingoid bases and of the N-acylated form (ceramides) are intimately linked. Structures and Occurrence In animal tissues, the most common or abundant of the sphingoid bases is sphingosine ((2S,3R,4E)-2-amino-4-octadecene-1,3-diol) or sphing-4E-enine, i.e., with a C18 aliphatic chain, hydroxyl groups in positions 1 and 3 and an amine group in position 2; the double bond in position 4 has the trans (or E) configuration. This was first characterized in 1947 by Professor Herbert Carter, who was also the first to propose the term “sphingolipides” for those lipids containing sphingosine. It is usually accompanied by the saturated analogue dihydrosphingosine (or sphinganine). Sphingoid bases are illustrated in Figure \(6\). For shorthand purposes, a nomenclature similar to that for fatty acids can be used; the chain length and number of double bonds are denoted in the same manner with the prefix 'd' or 't' to designate di- and trihydroxy bases, respectively. Thus, sphingosine is denoted as d18:1 and phytosphingosine is t18:0. The position of the double bond may be indicated by a superscript, i.e., 4-sphingenine is d18:1Δ4t or 4E-d18:1. While alternative nomenclatures are occasionally seen in publications, they are not recommended. The number of different long-chain bases that has been found in animals, plants and microorganisms now amounts to over one hundred, and many of these may occur in a single tissue or organism, but almost always as part of a complex lipid with an N-acyl-linked fatty acid and often phosphate or carbohydrate functional groups, as opposed to in the free form. The aliphatic chains can contain from 14 to as many as 28 carbon atoms, and most often they are saturated, monounsaturated or diunsaturated, with double bonds of either the cis or trans configuration. For example, the main dienoic long-chain base (sphingadienine) in human plasma is D-erythro-1,3-dihydroxy-2-amino-4-trans,14-cis-octadecadiene, and this is especially abundant in kidney, with more in women than in men. It is not present in zebra fish, widely used as a model species. Forms with three double bonds, such as sphinga-4E,8E,10E-trienine, sometimes with a methyl group in position 9, have been found the sphingolipids of some marine invertebrates and in a dinoflagellate. In addition, long-chain bases can have branched chains with methyl substituents in the omega‑1 (iso), omega‑2 (anteiso) or other positions, hydroxyl groups in positions 4, 5 or 6, ethoxy groups in position 3, and even a cyclopropane ring in the aliphatic chain in some organisms. N-Methyl, N,N-dimethyl and N,N,N-trimethyl derivatives of sphingoid bases have been detected in mouse brain. The main C18 components of long-chain bases of sphingomyelins of some animal tissues are accompanied by small amounts of C16 to C19 dihydroxy bases, although the latter attain higher proportions in tissues of ruminant animals. In gangliosides from human brain and intestinal tissues, eicosasphingosine (2S,3R,4E-d20:1) occurs in appreciable concentrations with variable amounts in different regions and membranes. However, human skin contains an especially wide range of isomers, including saturated, monoenoic and 6-hydroxy bases and phytosphingosines from C16 to C28 in chain-length. Shorter-chain bases are found in many insect species, and in the fruit fly, Drosophila melanogaster, which is widely used as a model species in genetic and metabolic experiments, the main components are C14 bases. In contrast to higher animals, nematodes such as Caenorhabditis elegans produce C17 iso-methyl-branched sphingoid bases, which are essential for normal sphingolipid function in the organism. The long-chain base composition of individual lipids can vary markedly between species, tissues, organelles and even different membranes within a single organelle. For example, the data in Table \(1\) is perhaps from an extreme example, but it illustrates that remarkable differences that can exist among lipids in one cellular component (rat liver mitochondria). Only part of the data from the paper cited is listed, but it illustrates that 3-keto-sphinganine, produced in the first step of sphingosine biosynthesis (see below) and normally a minor component of sphingolipids - often not detectable, can vary from 28 to 100% of the sphingoid bases depending on the lipid class and membrane within the organelle. Table \(1\): Long chain base composition of some lipid components of mitochondria from rat liver. Type Base (%) d18:1 d18:0-3keto t21:1 (phyto) Unidentified Ceramidesa 18 28 53 - Glucosylceramidesa 3 95 - 3 Lactosylceramidesb   100 a whole mitochondria; b mitochondrial inner membrane Data from Ardail, D. et al. FEBS Letts, 488, 160-164 (2001). Phytosphingosine or 4D-hydroxy-sphinganine ((2S,3R,4R)-2-amino-octadecanetriol) is a common long-chain base of mainly plant origin. It is a saturated C18-trihydroxy compound, although unsaturated analogues, for example with a trans (or occasionally a cis (Z)) double bond in position 8, i.e., dehydrophytosphingosine or 4D‑hydroxy-8-sphingenine, tend to be much more abundant. In many plant species, there are lipid class preferences also, and dihydroxy long-chain bases are more enriched in glucosylceramides than in glycosylinositolphosphoceramides, for example. This is true in the model plant Arabidopsis thaliana, where the data listed for whole tissue is probably representative largely of the latter lipid, as shown in Table \(2\) below. Table \(2\): Sphingolipid long-chain base composition of whole tissue and glucosylceramides from Arabidopsis thaliana. Base (%) t18:1 (8Z) t18:1 (8E) t18:0 d18:1 (8Z) d18:1 (8E) d18:0 Whole tissue 12 70 13   4 1 Glucosylceramides 44 22   5 28 2 Data from Sperling, P. et al. Plant Physiol. Biochem., 43, 1032-1038 (2005) Other plant long-chain bases have double bonds in position 4, which can be of either the cis or trans configuration, although trans-isomers are by far the more common, while the base d18:2Δ4E,8Z/E is relatively abundant in most plant species. In A. thaliana and related species, Δ4 long-chain bases are found mainly in the flowers and pollen and then exclusively as a component of the glucosylceramides. In general outwith Brassica species, the composition is dependent on species, but typically it is composed of up to eight different C18-sphingoid bases, with variable geometry of the double bond in position 8, i.e., (E/Z)-sphing-8-enine (d18:1Δ8), (4E,8E/Z)-sphinga-4,8-dienine (d18:2Δ4,8) and (8E/Z)-4-hydroxy-8-sphingenine (t18:1Δ8); d18:1Δ4, d18:0 and t18:0 tend to be present in small amounts only. Phytosphingosine is not restricted to plants, but is found in significant amounts in intestinal cells and skin of animals, with much smaller relative proportions in kidney. Although non-mammalian sphingoid bases in general tend to be poorly absorbed from the intestines, a small proportion of the phytosphingosine and related sphingoid bases found in animal tissues may enter via the food chain. Yeasts and fungi tend to have distinctive and characteristic long-chain base compositions. For example, filamentous fungi have 9-methyl-4E,8E-sphingadienine as the main sphingoid base in the glucosylceramides, as shown in Figure \(7\), but not in the ceramide phosphoinositol glycosides, while yeasts contain mainly the saturated C18 bases sphinganine and phytosphingosine, although some trans-4/8-unsaturated forms are usually present. Only a few bacterial species synthesize sphingolipids, but the family Bacteroidetes, which is abundant in the human gut is an important exception; they usually contain saturated (and branched) long-chain bases. Other pathogenic bacteria may utilize sphingolipids and sphingoid bases from their hosts. Sphingoid bases are surface-active amphiphiles with critical micellar concentrations of about 20 μM in aqueous solutions; they probably exist in the gel phase at physiological temperatures. In that they bear a small positive charge at neutral pH, they are unusual amongst lipids, although their pKa (9.1) is lower than in simple amines as a consequence of intra-molecular hydrogen bonding. Together with their relatively high solubility (> 1μM), this enables them to cross membranes or move between membranes with relative ease. In so doing, they increase the permeability of membranes to small solutes. In esterified form in complex lipids, they participate in the formation of ordered lipid domains in membranes such as rafts. In the complex sphingolipids, the sphingoid base is linked via the amine group to a fatty acid, including very-long-chain saturated or monoenoic and 2-hydroxy components, i.e., to form ceramides, which can be attached a polar head group, such as phosphate or a carbohydrate, via the primary hydroxyl moiety. An important exception is sphingosine-1-phosphate, which is not acylated and has signaling functions in cells akin to those of lysophospholipids. Biosynthesis and Metabolism Sphinganine biosynthesis The basic mechanism for the biosynthesis of sphinganine involves condensation of palmitoyl-coenzyme A with L-serine, catalyzed by the membrane-bound enzyme serine palmitoyltransferase, requiring pyridoxal 5’-phosphate as a cofactor, which binds to a specific lysine residue on the enzyme. The reaction occurs on the cytosolic side of the endoplasmic reticulum in animal, plant and yeast cells with formation of 3-keto-sphinganine as illustrated in Figure \(8\). This is believed to be the key regulatory or rate-limiting step in sphingolipid biosynthesis and is conserved in all organisms studied to date. Elimination of this enzyme is embryonically fatal in mammals and fruit flies. In mammals, serine palmitoyltransferase is a heterotrimer composed of two main subunits, designated SPTLC1 with either SPTLC2 or SPTLC3 (sometimes termed SPTLC2a and SPTLC2b, respectively). SPTLC1 is essential for activity, and it is ubiquitously expressed as is SPTLC2, while SPTLC3 is present in a relatively limited range of tissues and is most abundant in skin and placental tissue. In addition, there are two small subunits ssSPTA and ssSPTB (again other nomenclatures exist), which differ in a single amino acid residue, and may have regulatory functions; the active site is at the interface between the two main subunits. ssSPTA is essential for serine palmitoyltransferase function during development and hematopoiesis. A possible mechanism for the 1st step in the pathway, catalyzed by serine palmitotyltransferase, is shown in Figure \(9\). The addition of either of the two small subunits to the complexes changes the substrate preferences substantially and enables the synthesis of the wide range of homologs found in nature. In mammals, the SPTLC1-SPTLC2 complex forms C18 sphingoid bases specifically (with some C19, and C20), while the combination of SPTLC1 and SPTLC3 gives a broader product spectrum, including an anteiso-methylbranched-C18 isomer (from anteiso-methyl-palmitate as the precursor). Such branched bases are synthesized to a limited extent in human skin, but they are the main forms in lower invertebrates such as C. elegans. The activity of the serine palmitoyltransferase is governed by negative feedback and partly by orosomucoid (ORM-like or ORMDL) proteins, three in mammals (ORMDL1 to 3) and two in yeast (Orm1/2), which are ubiquitously expressed trans-membrane proteins located in the endoplasmic reticulum. The availability of serine is also an important factor. Figure \(10\) shows an interactive iCn3D model of the human serine palmitoyltransferase complex (7K0M). Figure \(10\): Human serine palmitoyltransferase complex (7K0M).. Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...tuGThTq9Vi4J58 • Gray and Plum (SPT1 A and E Chains • Light Blue and Blue (SPT2 B and F Chains) • Light Brown (small subunit A - ssA, C and G chains) • Yellow (ORM D and H) The second step in sphinganine biosynthesis is reduction of the keto group to a hydroxyl in an NADPH-dependent manner by a specific 3‑ketodihydrosphingosine reductase ('3KSR'), also on the cytosolic side of the endoplasmic reticulum, a step that must occur rapidly as the intermediate is rarely encountered in tissues. The enzymes are presumed to be in similar subcellular locations in plant cells. In plants, serine palmitoyltransferase is a heterodimer composed of LCB1 and LCB2 subunits with some homology to the mammalian enzymes, while in the yeast Saccharomyces cerevisiae, there are three subunits: Lcb1, Lcb2, and Tsc3. In the few bacteria that synthesize sphingoid bases, serine palmitoyltransferase is a water-soluble homodimer. The enzyme in the apicomplexan parasite Toxoplasma gondii is a homodimer also in contrast to other eukaryotes, but it is located in the endoplasmic reticulum. Free sphinganine formed in this way is rapidly N-acylated by acyl-coA to form dihydroceramides by dihydroceramide synthases, which in animals are located primarily on the endoplasmic reticulum, presumably on the cytoplasmic surface. Animals and plants have multiple isoforms of this enzyme, for which the abbreviated term ‘ceramide synthase’ is now widely applied as they utilize most other sphingoid bases, such as those produced by hydrolysis of sphingolipids, as substrates. They are unique gene products with each located on a different chromosome and with considerable variation in the expression of the enzymes in different cell types within each tissue. Each isoenzyme has distinct specificities for the chain-length of the fatty acyl-CoA moieties but to a limited extent only for the base, suggesting that ceramides containing different fatty acids have differing roles in cellular physiology. All of these enzymes have six membrane spanning regions, but the only substantial difference is in an 11-residue sequence in a loop between the last two putative transmembrane domains. Ceramides are central to all elements of sphingolipid biochemistry. These steps are illustrated in Figure \(11\). Humans and mice have six ceramide synthases, which utilize subsets of acyl-CoAs and thus producing ceramides with specific acyl chain lengths. Of these, ceramide synthase 2 is most abundant and is specific for coA esters of very-long-chain fatty acids (C20 to C26); it is most active in lung, liver and kidney. Ceramide synthase 1 is specific for 18:0 and is located mainly in brain with lower levels in skeletal muscle and testes. Ceramide synthase 3 is responsible for the unusual ceramides of skin and testes and uses C26-CoA and higher including polyunsaturated-CoAs with the latter tissue, while ceramide synthase 4 (skin, liver, heart, adipose tissue and leukocytes) uses C18 to C22-CoAs. Ceramide synthases 5 (lung epithelia and brain gray and white matter) generates C16 (mainly) and C18 ceramides, and ceramide synthase 6 (intestine, kidney and lymph nodes) produces C14 and C16 ceramides. However, hydroxylation and the presence or otherwise of double bonds in the acyl-coAs do not appear to influence the specificity of the ceramide synthases. Also, the expression of mRNA expression for ceramide synthases does not always correlate with the fatty acid composition of sphingolipids in a particular tissue, suggesting that other factors are involved in determining which molecular species are formed. One such is acyl-coenzyme A-binding protein (ACBP), which facilitates the synthesis of ceramides containing very-long fatty acids and stimulates ceramide synthases 2 and 3 especially. Insertion of the trans-double bond in position 4 to produce sphingosine occurs only after the sphinganine has been esterified in this way to form a ceramide as illustrated in Figure \(11\), with desaturation occurring at the cytosolic surface of the endoplasmic reticulum also. The desaturases were first characterized in plants, and this subsequently simplified the isolation of the appropriate enzymes in humans and other organisms. Two dihydroceramide desaturases have now been identified in animals and designated 'DEGS1 and DEGS2'. Both enzymes insert trans double bonds in position 4, but DEGS2 is a dual function enzyme that also acts as a hydroxylase to generate phytoceramides, i.e., to add a hydroxyl group on position 4. Distribution of the enzymes in tissues is very different, with DEGS1 expressed ubiquitously but highest in liver, Harderian gland, kidney and lung. DEGS2 expression is largely restricted to skin, intestine and kidney, where phytoceramides are more important. A considerable family of Δ4-sphingolipid desaturases has now been identified, and an early study by Stoffel and colleagues demonstrated that Δ4-desaturation involves first syn-removal of the C(4)- HR and then the C(5)-HS hydrogens. This appears to have been the first evidence that desaturases in general operate in this stepwise fashion. The enzyme responsible for the insertion of the cis-14 double bond into sphinga-4-trans,14-cis-dienine is the fatty acid desaturase 3 (FADS3), which utilizes ceramides containing sphingosine as the precursor. The only other known activity of this enzyme is to insert a cis-double bond in position 13 of the CoA ester of vaccenic acid (11t-18:1) to produce the conjugated diene 11t,13c-18:2. Synthesis of sphingoid bases de novo is essential in most organisms and inhibition of the biosynthetic pathways affects growth and viability. However, this can be tissue specific, as deletion of the liver-specific SPTLC2 in mice, was found to have no effect on liver function, while a comparable deletion of adipocyte-specific SPTLC1 caused major tissue defects. Presumably, the latter tissue is unable to take up enough sphingolipid from the circulation to remedy the problem. Deficiencies in SPTLC3 are related to dermal pathologies, and genetic variant of SPTLC3 are associated with dyslipidemia and atherosclerosis. The essentiality of sphingoid base synthesis in plants has been demonstrated in a similar manner in studies with mutants in which specific enzymes have been deleted. Phytosphingosine and plant ceramides: Phytosphingosine is formed from sphinganine, produced as above, by hydroxylation in position 4, possibly via the free base in plants, although it can be formed both from sphinganine and a ceramide substrate in yeasts. A single sphinganine C4‑hydroxylase is present in yeast, but Arabidopsis has two such enzymes (SBH1 and 2), which are critical for growth and viability. Much remains to be learned of the processes involved, but it is known that the enzyme responsible is closely related to a Δ4 desaturase. Indeed, it has been shown that there are bifunctional Δ4‑desaturase/Δ4-hydroxylases in Candida albicans and mammals, especially in keratinocytes (DEGS2 discussed above) with which either 4‑hydroxylation or Δ4‑desaturation is initiated by removal of the proR C-4 hydrogen. Sphinganine linked to ceramide is the substrate for 4-hydroxylation in intestinal cells. In Arabidopsis thaliana leaves, 90% of the sphingoid bases are phytosphingosine with a Δ8‑double bond. In plants in general, in addition to Δ4‑desaturation, two distinct types (20 gene products) of sphingoid Δ8-desaturase have been characterized that catalyse the introduction of a double bond at position 8,9 of phytosphingosine. These are evolutionarily distinct from the Δ4‑desaturases. One type produces the trans (E)-8 isomer mainly and the other mostly the cis (Z)-8 isomer, with overall the trans-isomer tending to predominate but dependent upon plant species. It appears that the trans isomer is formed when the hydrogen on carbon 8 is removed first, and the cis when carbon 9 is the point of attack. While the main group of Δ8-desaturases requires a 4‑hydroxysphinganine moiety as substrate, the second does not. In Arabidopsis, three different isoforms of ceramide synthase have been identified and denoted LOH1, LOH2 and LOH3. Phytosphingosine is used efficiently by LOH1 and LOH3 (class II synthases), but only LOH2 (class I synthase) uses sphinganine efficiently; LOH2 and 3 prefer unsaturated long-chain bases. Marked fatty acid specificity is also observed with LOH2 showing almost completely specific for palmitoyl-CoA and dihydroxy bases, while LOH1 shows greatest activity for 24:0- and 26:0-CoAs and trihydroxy bases; none utilize unsaturated acyl-CoA esters efficiently. In plants, fatty acid desaturases and hydroxylases are also closely related, and sphingolipid fatty acid α-hydroxylation is believed to occur on the ceramide, as opposed to the free acyl chain. It is believed that the Δ8‑desaturase utilizes ceramide as the substrate and the channels the products selectively into the synthesis of complex sphingolipids, while Δ4‑desaturation channels ceramides for synthesis of glucosylceramide. It has been established that long-chain bases with 4-hydroxyl groups are necessary for the viability of the filamentous fungus Aspergillus nidulans and for growth in plants such as A. thaliana. The presence of an 8E double bond confers aluminium tolerance to yeasts and plants, and it is important for chilling resistance in tomatoes. However, a trans-4 double bond in the sphingoid base does not appear to be essential for growth and development in Arabidopsis. Fungal sphingoid bases: Fungi produce trans Δ8-isomers only, but Δ4- and Δ8-desaturases do not occur in the widely studied yeast S. cerevisiae. In the biosynthesis of sphingoid bases in fungi, the double bonds in positions 4 and 8 and the methyl group in position 9 are inserted sequentially into the sphinganine portion of a ceramide, the last by means of an S-adenosylmethionine-dependent methyltransferase similar to plant and bacterial cyclopropane fatty acid synthases. In S. cerevisiae the ceramide synthase is a heteromeric protein complex, containing three subunits, Lag1, Lac1, and Lip1, of which the first two are homologous proteins that feature eight transmembrane domains. In the yeast Pichia pastoris, there is a distinct ceramide synthase, which utilizes dihydroxy sphingoid bases and C16/C18 acyl-coenzyme A as substrates to produce ceramides. The long-chain-base components of the ceramide are then desaturated in situ by a Δ4‑desaturase and the fatty acid components are hydroxylated in position 2. Further desaturation of the long-chain base component by a Δ8-(trans)- desaturase occurs before the methyl group in position 9 is introduced by an S-adenosylmethionine-dependent sphingolipid C-9 methyltransferase. As a final step a trans-double bond may be introduced into position 3 of the fatty acid component. These ceramides are used exclusively for the production of glucosylceramides, and it is believed that a separate ceramide synthase encoded by a different gene produces the ceramide precursors for ceramide phosphorylinositol mannosides. Viral sphingoid bases: The genome of an important marine virus (EhV) encodes for a novel serine palmitoyltransferase, which hijacks the metabolism of algal hosts to produce unusual hydroxylated C17 sphingoid bases; these accumulate in lytic cells of infected algae such as the important bloom-forming species Emiliania huxleyi. While this may seem a rather esoteric topic, viruses constitute a high proportion of the marine biome, and their control of the growth of algal blooms has global consequences. Unesterified sphingosine: A cycle of reactions occurs in tissues by which sphingoid bases are incorporated via ceramide intermediates into sphingolipids, which are utilized for innumerable functions, before being broken down again to their component parts. It is worth noting that all the free sphingosine in tissues must arise by this route, in particular by the action of ceramidases on ceramides. Five such ceramidases are known with differing pH optima and varying subcellular locations. The levels of free sphingoids and their capacities to function as lipid mediators, as shown in Figure \(12\), are controlled mainly by enzymic re‑acylation to form ceramides, although some is acted upon by sphingosine kinases to produce sphingosine-1-phosphate. Free sphingoid bases are absorbed by enterocytes following digestion of dietary sphingolipids in animals (including some from gut microorganisms), and while some of this is converted to complex sphingolipids, much is catabolized with the eventual formation of palmitic acid. Catabolism of sphingosine and other long-chain bases occurs after conversion to sphingosine-1-phosphate and analogues. In yeasts, an alternative means of detoxification has been reported in which an excess of phytosphingosine is first acetylated and then converted to a vinyl ether prior to export from the cells. Biological Functions of Unesterified Sphingoid Bases The primary function of sphingoid bases is to serve as a basic component of ceramides and complex sphingolipids, where variations in their compositions can influence the physical and biological properties of these lipids. Independently of this in their free (unesterified) form, they are important mediators of many cellular events even though they are present at low levels only in tissues (typically 25 and 50 nM in plasma), with intracellular levels determined by hydrolysis by ceramidases or by the action of sphingosine kinases (sphingosine-1-phosphate production). In animal cells, they inhibit protein kinase C indirectly, possibly by a mechanism involving interference with the binding of activators of the enzyme, such as diacylglycerols or phorbol esters. In addition, sphingoid bases are known to be potent inhibitors of cell growth, although they stimulate cell proliferation and DNA synthesis. They are involved in the process of apoptosis in a manner distinct from that of ceramides by binding to specific proteins and regulating their phosphorylation. While sphingosine does not appear to participate in raft formation in membranes, it may rigidify pre-existing gel domains in mixed bilayers, although any such effects will be dependent on local concentrations and pH. It should be noted that some of the biological effects observed experimentally may be due to conversion to sphingosine-1-phosphate. Free sphingosine has been implicated in various pathological conditions, and for example, plasma sphingosine levels are increased in hyperthyroidism and in patients with type 2 diabetes. Lysosomal storage of the lipid is an initiating factor in Niemann Pick type C disease, a neurodegenerative disorder, where it causes a change in calcium release leading to a buildup of cholesterol and sphingolipids. In the human adrenal cortex, sphingosine produced in situ by the acid ceramidase has a function in steroid production by serving as a ligand for steroidogenic factor 1 at the cell nucleus, which controls the transcription of genes involved in the conversion of cholesterol to steroid hormones. Unesterified sphingoid bases may have a protective role against cancer of the colon in humans. Thus, N,N‑dimethylsphingosine and dihydrosphingosine, like the deoxysphingoid bases, are known to induce cell death in a variety of different types of malignant cells. There is evidence that sphingadienes of plant and animal origin inhibit colorectal cancer in mouse models by reducing sphingosine-1-phosphate levels. In consequence, synthetic analogues of long-chain bases are being tested for their pharmaceutical properties. Free sphingosine is believed to have a signaling role in plants by controlling pH gradients across membranes. In addition, free long chain bases (and the balance with the 1-phosphate derivatives) are essential for the regulation of apoptosis in plants. Ceramides Structure and Occurrence The structure of ceramide is shown again in Figure \(13\). Figure \(13\): Structure of ceramides (with varying fatty acids in ester link) Ceramides consist of a long-chain or sphingoid base linked to a fatty acid via an amide bond. They are essential intermediates in the biosynthesis and metabolism of all sphingolipids including the complex sphingolipids in which the terminal primary hydroxyl group is linked to carbohydrate, phosphate, and so forth (sphingomyelin, glycosphingolipids and gangliosides) as shown in Figure \(14\). They are also the primary source of unesterified sphingoid bases and of the important biological mediators sphingosine-1-phosphate and ceramide-1-phosphate. At the last count, 33 different enzymes were known to participate in ceramide metabolism. While ceramides are rarely found as such at greater than trace amounts in tissues other than skin, they can exert important biological effects of their own at these low levels. They are present in membranes where they participate in the formation of raft domains. Each organism and indeed each tissue may synthesize ceramides in which there are a variety of di- and trihydroxy long-chain bases linked to fatty acids. As discussed previously, the fatty acids consisting mainly of longer-chain (up to C24 or greater) saturated and monoenoic (mainly (n-9)) components, sometimes with a hydroxyl group in position 2. Other than in certain testicular cells, polyunsaturated fatty acids do not occur. More than 200 structurally distinct molecular species of ceramides have been characterized from mammalian cells. In plants, 2-hydroxy acids predominate sometimes accompanied by small amounts of 2,3-dihydroxy acids. Although small amounts of free ceramides are produced in all tissues as required for the specific biological functions described below, most is converted rapidly to more complex sphingolipids, including sphingomyelin (in animals) and the various glycosylceramides. The ceramides in skin are a remarkable exception to this rule, and as such they are discussed separately below. A shorthand nomenclature simply combines those used conventionally for fatty acids and long-chain bases to denote molecular species of ceramides, including those as components of more complex lipids, e.g. N-palmitoyl-sphingosine is d18:1-16:0. Ceramides containing sphinganine are sometimes termed ‘dihydroceramides’. Ceramide Biosynthesis Ceramide production is complex and involves at least three pathways. Biosynthesis de novo takes place in the endoplasmic reticulum with palmitoyl-CoA and serine as the precursors for the long-chain base component, which is subsequently converted to ceramide. Biosynthesis of the very specific fatty acids in ceramides involving various chain elongases (ELOVL) requires consideration also. Alternative routes for ceramide production involve regeneration from complex sphingolipids. For example, in animals in the sphingomyelinase pathway, conversion of sphingomyelin into ceramides (and vice versa) occurs in the plasma membrane, Golgi and mitochondria. Finally, the polar moieties of complex glycosphingolipids can be removed by various hydrolytic enzymes in the lysosomal compartment to recover the ceramides (or their component parts) in a re-cycling/catabolic process. As these biosynthetic or metabolic pathways are located in different organelles, specific pools of ceramide and sphingolipids result with differing biological properties and functions. Ceramide synthesis de novo: The first of these pathways is described in mechanistic. In brief in animals, sphinganine is coupled to a long-chain fatty acid to form dihydroceramide by means of one of six ceramide synthases in the endoplasmic reticulum mainly, before the double bond is introduced into position 4 of the sphingoid base. Of these, ceramide synthase 2 is most abundant and is specific for CoA esters of very-long-chain fatty acids (C20 to C26); it is most active in the central nervous system. Ceramide synthase 1 is specific for 18:0 and is located exclusively in brain and skeletal muscle, ceramide synthases 5 and 6 generate 16:0-containing ceramides, and ceramide synthase 3 is responsible for the unusual ceramides of skin and testes. Figure \(15\) shows again the synthesis of ceramide from sphinganine and palmitoyl-CoA (a repeat of Figure \(11\) Each synthase has six membrane-spanning domains and contains a characteristic motif with the specific structures required for catalysis and substrate binding that are essential for its activity, and they have been shown to differ primarily in an 11-residue sequence in a loop between the last two putative transmembrane domains. In addition to separate transcriptional regulation of each of these enzymes, ceramide synthase activity is modulated by many different factors including reversible dimerization, while ceramide synthase 2 has a sphingosine-1-phosphate binding motif and this lipid may inhibits its activity. Acyl-coenzyme A-binding protein (ACBP) facilitates the synthesis of ceramides containing very-long fatty acids and stimulates ceramide synthases 2 and 3 especially. Most of the ceramides generated in this way are rapidly utilized for synthesis of complex sphingolipids, especially sphingomyelin and hexosylceramides, to ensure that cellular ceramide concentrations are regulated to control their biological activities. In mammalian cells, most complex glycerolipids are synthesized in the endoplasmic reticulum prior to their transport to their final subcellular locations, but the process is rather different for sphingolipids. Ceramide is synthesized on the cytoplasmic leaflet of the endoplasmic reticulum, but subsequent formation of complex sphingolipids occurs in the Golgi apparatus, and a key cytoplasmic protein, ceramide transporter or 'CERT' (CERamide Trafficking), mediates the transport of ceramide between these organelles in a non-vesicular manner. It has a number of distinct functional domains, including an N-terminal phosphatidylinositol-4-monophosphate (PI(4)P)-binding or Pleckstrin homology (PH) domain, which targets the Golgi apparatus, and a C-terminal ‘START’ domain, which can recognize ceramide species with the natural D-erythro stereochemistry, including dihydroceramide and phytoceramide (but not sphingosine), and holds them within in a long amphiphilic cavity by hydrogen bonding with all three polar atoms of the sphingoid motif. There is also a short peptide motif (FFAT) that recognizes a specific protein in the endoplasmic reticulum. There is sufficient flexibility in the body of the protein to enable transfer of ceramide from the endoplasmic reticulum to the Golgi without free movement through the cytosol. Very-long-chain ceramides containing 24:0 or 24:1 fatty acids turn over much more rapidly in animal cells than those containing 16:0 or 18:0 fatty acids, because of the more rapid conversion of the former into complex sphingolipids, where they may regulate the levels and perhaps the biological functions of the latter. In contrast, ceramides containing d16:1 and d18:1 sphingoid bases turnover at similar rates so do not affect the flux of ceramides through these pathways. The CERT protein is a major factor in this specificity, as it extracts ceramides from membrane bilayers with a preference for those required for synthesis of complex sphingolipids. Removal of ceramide by this process provides the gradient that enables the process to continue, and prevents an accumulation of ceramide in the endoplasmic reticulum that might otherwise be disruptive to the membrane and even cause cell death. While the transfer process itself is not dependent on ATP, the overall process requires ATP, possibly to keep PI(4)P in a phosphorylated form, and the multiple factors that control the biosynthesis of this lipid must also influence sphingolipid metabolism. As a neutral lipid, ceramide can flip readily across membrane leaflets, and this is also necessary for the synthesis of sphingomyelin, which occurs on the lumen of the Golgi. The pool of ceramide utilized for the synthesis of glycosylceramide is delivered to the Golgi by a separate transport mechanism that also does not require ATP. In addition, some ceramide synthesis occurs in mitochondria although this has the potential to lead to cell death. Regulation of ceramide and subsequent sphingolipid biosynthesis is crucial as an excess of sphingolipids can be toxic, while reduced synthesis can inhibit cell proliferation. Some ceramides are transported from the liver to other tissues in plasma lipoproteins, but especially subclasses HDL2 and HDL3, i.e. those containing apolipoprotein B. There is a suggestion that transport of ceramides via lipoproteins could be a paracrine mechanism to regulate the metabolism of other cells. Ceramides are also produced during the catabolism of other complex sphingolipids, and especially by the action of one or other of the sphingomyelinases or of phospholipase C on sphingomyelin in animal tissues as part of the 'sphingomyelin cycle' as shown in Figure \(16\). Many agonists including chemotherapeutic agents, tumor necrosis factor-alpha, 1,25-dihydroxy-vitamin D3, endotoxin, gamma-interferon, interleukins, nerve growth factor, ionizing radiation and heat stimulate hydrolysis of sphingomyelin to produce ceramide. In addition, reversal of the sphingomyelin synthesis reaction may generate ceramide, and some may be produced by operation of the enzyme ceramidase in reverse (see next section). Such reactions are much more rapid than synthesis de novo, so they are of special relevance in relation to the signaling functions of ceramides, especially when they occur at the plasma membrane. For example, in this context, the acid sphingomyelinase may be especially important by generating the ceramides that initiate the train of events that leads to apoptosis (see below). Glycosphingolipids can be hydrolyzed by glycosidases to ceramides also in tissues, but the process tends to be less important in quantitative terms (other than in skin). The key enzymes of sphingolipid metabolism were first characterized from the yeast Saccharomyces cerevisiae, and these were found to be sufficiently similar to the corresponding enzymes in mammals to facilitate their study in the latter. As discussed above, there are specific ceramide synthases that utilize specific fatty acids for ceramide biosynthesis in animals, and knowledge is slowly being acquired of how these are compartmentalized and regulated within cells. Thus, the synthesis and subsequent catabolism of ceramides involves a complex web of at least 28 distinct enzymes, including six ceramide synthases and five sphingomyelinases, which are all products of different genes. Each of these enzymes may produce distinctive molecular species of ceramides with their own characteristic biological properties. It has been determined that ceramide species containing very-long-chain fatty acids (C24) turnover more rapidly than those containing C16/18 components. Ceramide Catabolism In animals, ceramide metabolism and function are controlled in part by the action of ceramidases, which cause hydrolysis forming sphingoid bases and free fatty acids, and indeed this is the only route to the formation of unesterified sphingosine. This is illustrated in Figure \(17\). Five such enzymes are known in humans, classified according to their pH optima, i.e. acid (‘ASAH1’), neutral (‘ASAH2’, which differs between humans and animals), and alkaline (three enzymes - ‘ACER1 to ACER3’), with differing cellular locations and fatty acid specificities and with the potential to affect distinct signaling and metabolic events. The acid ceramidase is of particular importance, and aberrations in its synthesis or activity is involved in several human disease states, including the rare autosomal-recessive Farber disease where there is a deficiency in the enzyme so ceramide accumulates; ceramide containing 26:0 in the blood is considered to be a biomarker for diagnosis of the disease. ASAH1 is located in the lysosomes and hydrolyses ceramides with small to medium-chain fatty acid components (C6 to C18) most efficiently. The neutral ceramidase is located in the plasma membrane and Golgi, especially of intestinal epithelial cells and colorectal tissues, and prefers long-chain components (C16 to C18); it also catalyzes the reverse reaction, and this may be a means of ceramide synthesis in mitochondria. ACER1 and ACER2 are found in the endoplasmic reticulum and Golgi, respectively, and they prefer species with very-long-chain acyl groups. ACER3 is present in both the endoplasmic reticulum and Golgi; it has a marked specificity for ceramides, dihydroceramides, and phytoceramides linked to unsaturated long-chain fatty acids (18:1, 20:1 or 20:4) in vitro at least. Neutral/alkaline ceramidase activity has also been found in mitochondria and nuclei. In Arabidopsis, an alkaline ceramidase (AtACER) can hydrolyze phytosphingosine-containing ceramides, and a related enzyme from rice has a preference for d18:1Δ4-ceramide; the latter can function in reverse to increase the content of C26- and C28-phytoceramides. Several neutral ceramidases (AtNCERs) have been identified, but there does not appear to be an equivalent to the acid ceramidase in plants. Ceramidases are also present in lower organisms such as Pseudomonas aeruginosa and slime molds, where they are secreted proteins rather than integral membrane enzymes. A neutral ceramidase only is found in prokaryotes, including some pathogenic bacteria. Sphingoid bases released by the action of acid ceramidase can escape from the lysosomes and be re-utilized for ceramide biosynthesis through the action of a ceramide synthase. This has been termed the ‘salvage’ pathway and is important in both quantitative and biological terms. For example, it has been estimated that it contributes from 50 to 90% of sphingolipid biosynthesis. The biological functions of ceramides are discussed below, but there are reasons to believe that ceramides derived from the salvage pathway are spacially and thence functionally distinct from those synthesized de novo. In addition, sphingoid bases released in this way have their own biological functions, which includes utilization for the synthesis of the biologically important metabolite sphingosine-1-phosphate. Therefore, regulation of ceramidase action is central to innumerable biological processes in animals. Biological Functions of Ceramides The role of ceramides in the biosynthesis of complex glyco- and phospho-sphingolipids are discussed elsewhere in this text. Ceramides, like other lipid second messengers in signal transduction, are produced rapidly and transiently in response to specific stimuli in order to target specific proteins, for example to activate certain serine/threonine protein kinases or phosphatases. They may also regulate cellular processes by influencing membrane properties. While they can be produced by synthesis de novo for such functions, activation of one of the sphingomyelinases under physiological stress or other agents is a more rapid means of generation in animal tissues at least. In fact, ceramides appear to be formed under all conditions of cellular stress by a multiplicity of activators in eukaryotic organisms. However, it should be noted that ceramides with different fatty acid and long-chain base (molecular species) compositions are formed in different compartments or membranes of the cell by various mechanisms over different time scales and potentially with distinct functions. The biological functions of those ceramides containing medium-chain (up to C14), long-chain (C16 and C18), and very-long-chain (C20 and longer) fatty acids, in particular, may have to be considered separately. Physical properties: Unsaturation in the sphingoid backbone augments intramolecular hydrogen bonding in the polar region, which permits a close packing of ceramide molecules and a tight intramolecular interaction in membranes. A further important factor in this context is the length of the fatty acyl moiety, as shorter-chain ceramides tend to produce a positive curvature in a lipid monolayer, while long-chain molecules have the opposite effect and possess a marked intrinsic negative curvature that facilitates the formation of inverted hexagonal phases as well as increasing the order of the acyl chains in bilayers. By their interactions with ion channels, ceramides influence the permeability of membranes and render bilayers and cell membranes permeable to solutes that vary from small- up to protein-size molecules. While ceramides are minor components of membranes in general, their physical properties ensure that they are concentrated preferentially into lateral liquid-ordered microdomains (a distinct form of 'raft' termed ‘ceramide-rich platforms’), although these effects are again chain-length specific. These domains differ appreciably in composition from those rafts enriched in sphingomyelin and cholesterol, and ceramides containing C12 to C18 fatty acids can in fact displace cholesterol from rafts to modify their physical properties. Ceramides are generated within rafts by the action of acid sphingomyelinase, causing small rafts to merge into larger units and modifying the membrane structure in a manner that is believed to permit oligomerization of specific proteins such as cytokines and death receptors. Ceramides are also essential for the formation and/or secretion of exosomes by facilitating or inducing membrane curvature. In contrast, sphingosine, sphingosine-1-phosphate and ceramide-1-phosphate do not facilitate raft formation. Through the medium of these modified rafts, ceramides are able to function in signal transduction. Specific receptor molecules and signaling proteins are recruited and cluster within such domains, thereby excluding potential inhibitory signals, while initiating and greatly amplifying primary signals. It is believed that ceramide-rich platforms amplify both receptor- and stress-mediated signaling events and thence may influence various disease states. Ceramide-enriched membrane domains formed in response to sphingomyelinase activity are sites for endocytic uptake of pathogens because of a concentration of pathogen receptors and signaling complexes, and in particular these can enhance viral infections, including Norovirus, Japanese encephalitis virus, Ebola and possibly SARS-CoV-2. However, elevated levels of ceramide inhibit cellular uptake of the HIV virus. Although ceramides and diacylglycerols have structural similarities, their occurrence, location, and behavior in membranes are different. Ceramides cross synthetic lipid bilayers relatively quickly in vitro, but it is not clear whether they can flip across more complex biological membranes equally readily, especially in the ceramide-rich platforms. Restricted flipping could have important effects on the signaling role of ceramides in that those generated by different enzymes on each side of a membrane could have distinct functions. Enzyme activation: In general, ceramides tend to modify intracellular signaling pathways to slow anabolism and promote catabolism. Amongst a wide range of biological functions in relation to cellular signaling, ceramides are especially important in triggering apoptosis, and they have also been implicated in the activation of various protein kinase cascades, dependent on the site of generation. The mechanism of these interactions is the subject of intensive study at present, but in relation to the latter, two intracellular targets for ceramide action of special importance have been discovered – at least two protein phosphatases (ceramide-activated protein phosphatases) and a family of protein kinases (ceramide-activated protein kinases). For example, the phosphatase may be involved in the regulation of glycogen synthesis, insulin resistance, and response to apoptotic stimuli. Ceramides generated by the action of sphingomyelinase and by synthesis de novo are both important to the process, while ceramidases have contrasting effects in these and other biological effects of ceramides. Apoptosis: The role of ceramides in the regulation of apoptosis, and cell differentiation, transformation, and proliferation has received special attention. Apoptosis is a normal process, which occurs in response to oxidative stress in particular, in which a cell can be considered to actively ‘commit suicide’. It is essential for many aspects of normal development and is required for maintaining tissue homeostasis. There are two pathways - 'extrinsic' initiated in the plasma membrane by ligation of so-called 'death factors', such as the tumor necrosis factor-α (TNF-α), and 'intrinsic' induced by external actions in mitochondria, e.g. by DNA damage, oxidation or radiation injury. Although the mechanism of the ceramide interaction with these pathways is uncertain, it is clear that a cascade of reactions is initiated that culminates in the release of intracellular proteases of the caspase family to promote apoptosis. In dysfunctional mitochondria, one mechanism involves the formation of channels in the membrane that enable the release of specific mitochondrial proteins that include caspases. Ceramides with fatty acids of differing chain lengths are believed to function in different ways, and 16:0-ceramide generated by ceramide synthase 6 is especially pro-apoptotic, for example, while ceramides with very-long-chain fatty acids accumulate in necroptosis, a form of apoptosis. On the other hand, ceramides containing 2-hydroxy acids in keratinocytes appear to be protective against apoptosis. Ceramides induce the related process of cellular senescence also. Failure to properly regulate apoptosis can have catastrophic consequences, and many disease states, including cancer, diabetes, neuropathies, Alzheimer's disease, Parkinson's disease, and atherosclerosis, are thought to arise from the deregulation of apoptosis. For example, ceramides have been implicated in the actions of TNF-α and in the cytotoxic responses to amyloid Aβ peptide, which are involved in Alzheimer’s disease and neurodegeneration. In addition, ceramides appear to be involved in many aspects of the biology of aging and of male and female fertility. These effects may hold implications for diseases associated with obesity and insulin resistance, including again diabetes and cardiovascular disease. Similarly, ceramides are intimately involved in the induction of autophagy, the 'maintenance' process by which cellular proteins and excess or damaged organelles are removed from cells by engulfing them in a membrane-enclosed cellular compartment called the phagosome. In particular, maturing phagosomes are enriched in very-long-chain ceramides. While this process is beneficial in that it aids the recycling of cellular nutrients, the presence of excess ceramide can lead to unnecessary apoptosis. As animals and plants have multiple isoforms of ceramide synthase that are specific for the chain length of the base and fatty acid, it has been suggested that ceramides containing different fatty acids have distinct roles in cellular physiology. In particular, C16 ceramide appears to be especially important in apoptosis in non-neuronal tissues, while C18 ceramide has growth-arresting properties and may be involved in apoptosis in some carcinomas treated with chemotherapy agents. In addition, a transferase has been identified that transfers the acetyl group from platelet-activating factor to sphingosine with a high specificity. The product, N-acetylsphingosine - the simplest of all ceramide molecules, has signaling functions that are distinct from those of the parent lipids or of other ceramides; it does not enter the salvage pathway in cancer cells in vitro and is cytotoxic. In contrast, the ceramide metabolite, sphingosine-1-phosphate, has opposing effects on cell survival and proliferation. As ceramide and sphingosine-1-phosphate are inter-convertible via sphingosine as an intermediate, which also has pro-apoptopic activity, the balance between these lipids and with ceramide-1-phosphate is obviously of great metabolic importance. It has been termed the ‘sphingolipid-rheostat’, as illustrated in Figure \(18\). Plants: Comparatively little information is available on the role of ceramides in cell signaling in plants, but there are suggestions that sphingolipid catabolic products may be linked to programmed cell death, signal transduction, membrane stability, host-pathogen interactions, and stress responses. For example, there is evidence that enhanced synthesis of ceramides with very-long-chain fatty acids and trihydroxy sphingoid bases by ceramide synthases LOH1 and LOH3 promotes cell division and growth, while in contrast, accumulation of the ceramide species C16 fatty acid with a dihydroxy sphingoid base, due to LOH2 overexpression, leads to plant dwarfing and programmed cell death. Ceramides aggregate in rafts in plant membranes, together with other sphingolipids and sterols, as in animal tissues. Similarly, in the yeast S. cerevisiae, widely used as a model organism, it has been reported that ceramide species with different N-acyl chains and sphingoid bases are involved in the regulation of different sets of functionally related genes. Skin Ceramides The mammalian skin forms the protective barrier between the internal tissues of the host and the hostile external environment, which can include chemicals, ultraviolet light, mechanical damage, and pathogenic microorganisms, while preventing the loss of water and electrolytes. It consists of stratified layers of increasingly differentiated cells or keratinocytes of which the basal layer is responsible for the renewal of the tissue but begins to migrate upwards and differentiate, while accumulating specific lipids and proteins that change the cellular architecture. Eventually, the keratinocytes lose their nucleus and become flattened structures of insoluble protein surrounded by lipids termed ‘corneocytes’ in the outermost impermeable layer or stratum corneum. By secreting peptides and proteins that possess antimicrobial activity, keratinocytes add to the defensive capability of skin against commensal microorganisms and opportunistic pathogens, and this is reinforced by lipid mediators such as free sphingoid bases and eicosanoids in the stratum corneum and free fatty acids in sebum. The stratum corneum contains high levels of ceramides (as much as 50% of the total lipids), including O-acylceramides, which exist both in the free form and linked by ester bonds to structural proteins. They are present mainly in the extracellular domains (interstices) and are accompanied by nearly equimolar amounts of cholesterol and free fatty acids, a ratio that is believed to be essential for the normal organization of the tissue into the membrane structures that are responsible for the functioning of the epidermal barrier. In contrast to other biological membranes, the lipid organization in the membranes of skin consists of two lamellar phases, which form crystalline lateral phases mainly, with repeat distances of approximately 6 and 13 nm. Small sub-domains of lipids in a liquid phase may also exist. Some of these skin ceramides have distinctive structures not seen in other tissues, and many different forms are commonly recognized. They can contain the normal range of longer-chain fatty acids (a), e.g. formula 1 in the figure, some with hydroxyl groups in position 2 (a*), e.g. formula 2, linked both to dihydroxy bases with trans-double bonds in position 4 or to trihydroxy bases. This is illustrated in Figure \(18\). In addition, there are O‑acyl ceramides in which a unique very-long-chain fatty acid component (typically C30 or C32) has a terminal hydroxyl group, and this may be in the free form or esterified with linoleate (c), e.g., formulae 3 and 4; the sphingoid base can be either di- (b) or trihydroxy (b*), e.g., formula 4; the latter is not a common feature in sphingolipids of animal origin, and can include both phytosphingosine and the unique 6‑hydroxy-4-sphingenine in human epidermis. Ceramides of type 1 in which the 1-O-hydroxyl group of the sphingoid base is acylated by a very-long-chain fatty acid are also present (1‑O‑acylceramides - illustrated above); these comprise 5% of the total ceramides in the epidermis of mice and humans and comprise as much as 700 molecular species. In all, 15 classes of free ceramides and 3 classes of covalently bound ceramides with up to 1700 distinct molecular species have been identified. Such lipids were first studied in detail in the skin of the pig as a convenient experimental model, but they have been characterized in humans and rats. In addition, several molecular forms of glucosylceramide, based on similar ceramide structures, have been characterized in skin, and these are also essential for its proper function. Depending on the particular layer of the skin (keratinocytes, stratum corneum, etc.), the lipid composition can vary. These lipids have an obvious role in the barrier properties of the skin, limiting the loss of water and solutes and at the same time preventing the ingress of harmful substances. As the aliphatic chains in the ceramides and the fatty acids are mainly non-branched long-chain saturated compounds with a high melting point and a small polar head group, the lipid chains are mostly in a solid crystalline or gel state, which exhibits low lateral diffusional properties and low permeability at physiological temperatures. There is a report that the stratum corneum layer of the skin has a water permeability only one-thousandth that of other biomembranes, for example. Natural and synthetic ceramides are now commonly added to cosmetics and other skin care preparations. Most steps in the biosynthesis of ceramides linked to ω-O-acylated fatty acids occur in the endoplasmic reticulum of keratinocytes. First, fatty acid synthesis of very-long-chain (and ultra-long-chain, ≥C26) acyl-CoA de novo must take place, requiring the chain-elongation enzymes ELOVL1 and ELOVL4. Desaturation can occur, and importantly oxidation in the 2 (α) and terminal (ω) positions. The ω‑hydroxylation step requires an enzyme of the cytochrome P450 family, designated CYP4F22, of the kind involved in the synthesis of hydroxy-eicosatetraenoic acids (HETE). Mutations are a cause of lamellar ichthyosis, and knockout mice deficient in the equivalent enzyme were found to die within 8 hours of birth. Ceramides are first synthesized by ceramide synthase 3 (CERS3), which has a high specificity for very-long-chain fatty acids (>C26) with the incorporation of the ω‑hydroxy fatty acid. This is acylated with linoleate by the action of an unusual enzyme related to the phospholipase A family, PNPLA1, which catalyzes esterification by first releasing linoleate from triacylglycerols in the skin while acting as an acyltransferase to link the linoleate directly to the ω-hydroxyl moiety of the ultra-long chain fatty acid. PNPLA1 is unique among phospholipases in that it is involved in the metabolism of sphingolipids rather than glycerophospholipids and catalyzes transacylation rather than hydrolysis. In addition, some linoleate for this purpose is released from triacylglycerols by the action of the adipose tissue lipase aided by a protein ABHD5. This process is vital for proper skin barrier function and keratinocyte differentiation, as mice with defective triacylglycerol biosynthesis and metabolism, including a deficiency of the acyl-CoA synthase ACSL1, are unable to synthesis ω‑O‑acylceramides and have an impaired skin barrier. Mutations in the human PNPLA1 gene are believed to be the cause of autosomal recessive disease congenital ichthyosis. The resulting ceramides are converted to the complex sphingolipids sphingomyelin and especially glucosylceramide, which are transferred with the aid of ATP-binding cassette (ABC) transporters together with degradative enzymes into the stratum corneum via specific organelles termed 'lamellar bodies.' These organelles must fuse with the apical plasma membrane of the outermost cell layer of the epidermis in order that their contents can be secreted. It is only then that the final step of hydrolysis of the lipid precursors occurs in the extracellular spaces of the stratum corneum, i.e. ceramides are generated from sphingomyelin by the action of acid sphingomyelinase and from glucosylceramides by β-glucocerebrosidase. This mechanism ensures that ceramides, with their potentially harmful biological activities, never accumulate within nucleated cells. Eventually, ceramides with a terminal ω-hydroxyl group in the fatty acyl moiety are bound covalently to the proteins of the cornified envelope, especially to involucrin. This is illustrated in Figure \(19\). Sphingomyelin and Related Sphingophospholipids Structure and Occurrence of Sphingomyelin Sphingomyelin or ceramide 1-phosphocholine consists of a ceramide unit with a phosphorylcholine moiety attached to position 1 of the sphingoid base component. It is thus the sphingolipid analog of phosphatidylcholine, and like that lipid it is zwitterionic. The d18:1/16:0 molecular species is illustrated as an example in Figure \(20\). Sphingomyelin is primarily of animal origin and is a ubiquitous component of all animal cell membranes, from mammals to nematodes (and in a few protozoa), where it is by far the most abundant sphingolipid. Indeed, it can comprise as much as 50% or more of the lipids in certain tissues, though it is usually lower in concentration than phosphatidylcholine. For example, it makes up about 10% of the lipids of the brain, where it is a key constituent of myelin, but 70% of the phospholipids of the human lens. Like phosphatidylcholine, sphingomyelin tends to be in greatest concentration in the plasma membrane of cells (up to 20%), and in the endocytic recycling compartment and trans Golgi network. It is also abundant in the nucleus where it is the main phospholipid associated with chromatin, but there is very little in the endoplasmic reticulum (2 to 4%) and even less in mitochondria. All the sphingomyelin in human erythrocyte membranes is in the outer leaflet, and ~90% of that in the plasma membrane of nucleated cells is in the outer leaflet. All lipoprotein fractions in plasma contain appreciable amounts of sphingomyelin with a higher proportion in the VLDL/LDL. Sphingomyelin is the single most abundant lipid in erythrocytes of most ruminant animals, where it replaces phosphatidylcholine entirely. In this instance, there is known to be a highly active phospholipase A that breaks down the glycerophospholipids, but not sphingomyelin. Sphingomyelin is not synthesized in plants or fungi, which produce the sphingophospholipid ceramide phosphoinositol and related lipids instead, or in bacteria, and its evolutionary significance is a matter for speculation. However, a number of bacteria and viruses utilize sphingomyelin or its metabolism in their hosts for growth and viability. Sphingosine is usually the most abundant long-chain base constituent, together with sphinganine and C20 homologues, although other bases can be present, especially in ruminant animals. In contrast, sphinganine is the major sphingoid base in the sphingomyelin of human lens membranes, linked mainly to 16:0. Typically, the fatty acids are very-long-chain saturated and monounsaturated, including odd-numbered components. In comparison to the glycosphingolipids, 2‑hydroxy acids are only rarely detected and then in small amounts, but they are found in testes, spermatozoa, kidney and skin sphingomyelin, for example. The absolute proportions of each fatty acid and sphingoid base can vary markedly between tissues and species, and some of the variability in compositions can be seen from the data in Table \(3\) and Table \(4\) below. Table \(3\): Fatty acid compositions of sphingomyelin (wt % of the total) in some animal tissues. Source Fatty acids 16:0 18:0 18:1 20:0 22:0 22:1 23:0 23:1 24:0 24:1 Egg 66 10 1 4 6 1 2 - 5 3 Bovine brain 3 42 - 6 7 3 3 3 6 27 Cow's milk 14 3 1 1 22 - 32 - 19 5 Adapted from Ramstedt, B. et al. Analysis of natural and synthetic sphingomyelins using high-performance thin-layer chromatography. Eur. J. Biochem., 266, 997-1002 (1999); DOI. Table \(4\): Long-chain base compositions of sphingomyelin (wt % of the total) in some animal tissues. Source Sphingoid base d16:0* d17:0 d17:1 d17:1-methyl d18:0 d18:1 d19:0 Egg         7 93 Bovine brain         19 81 Cow's milk 9 15 8 11 10 44 3 Also from Ramstedt, B. et al. Eur. J. Biochem., 266, 997-1002 (1999); DOI. * d = dihydroxy base Palmitic acid (16:0) is the most common fatty acid component of sphingomyelin in peripheral cells of mammals, while stearic acid (18:0) is more abundant in neural tissue, but this only hints at the potential complexity as there can be variability within tissues. For example, about 60% of the fatty acids of the sphingomyelin of the grey matter of the human brain consist of stearic acid (18:0), while lignoceric (24:0) and nervonic (24:1) acids make up 60% of the corresponding lipid of white matter, although this is dependent on the stage of development. During the first two years of life, the 18:0 concentration in sphingomyelin of white matter decreases from 82% to 33%, while the proportions of 24:0 and 24:1 increase. This pronounced shift from long-chain to very-long-chain sphingomyelins is not observed in the cerebral cortex. Approximately 100 molecular species of sphingomyelin have been detected in human plasma. Although polyunsaturated fatty acids such as arachidonic acid are rarely present, they have sometimes been mistakenly identified in the literature. Exceptions are the sphingomyelins of testes and spermatozoa, which contain very-long-chain polyunsaturated fatty acids (up to 34 carbon atoms), the major components being 28:4(n-6) and 30:5(n-6) with a proportion having hydroxyl groups in position 2. Biosynthesis, Metabolism and Function of Sphingomyelin The biosynthesis of sphingomyelin is distinct from that of phosphatidylcholine and indeed depends upon it, as it involves the transfer of phosphorylcholine from phosphatidylcholine to ceramide, synthesized in the endoplasmic reticulum, with the liberation of 1,2-diacyl-sn-glycerols. as illustrated in Figure \(21\). The reaction is catalyzed by a ceramide choline-phosphotransferase (sphingomyelin synthase or SMS) and takes place primarily on the luminal side of the trans-Golgi but also in the plasma membrane, with two related enzymes each with six transmembrane domains and their N- and C-termini facing the cytosol, i.e., SMS1 and SMS2. Both enzymes are present in the Golgi, but only SMS2 is in the plasma membrane (facing the extra-cellular space in this instance) and may be necessary for the formation of raft domains (see below). SMS2 is also present in the membranes of nuclei from rat liver cells. It is noteworthy that in the absence of ceramide, both SMS1 and 2 have phospholipase C activity, and so may regulate the steady-state levels of phosphatidylcholine and diacylglycerols as well as that of sphingomyelin. The reaction does not use free phosphorylcholine or CDP-choline as a donor. Figure \(22\) shows an interactive iCn3D model of the AlphaFold model of human Golgi membrane phosphatidylcholine:ceramide cholinephosphotransferase 1, also called sphingomyelin synthase 1 (Q86VZ5). Figure \(22\): . Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...v6s5wHRgPXKJC6 The gray helices are the transmembrane helices. A cytoplasmic protein:protein interaction domain called SAM (sterile alpha motif) is shown in magenta. The other cytoplasmic C-terminal domain is shown in cyan Obviously much of the cytoplasmic domain is disordered in this computational structure. Side chains involved in binding phosphatidylcholine are shown as sticks colored CPK. Note that two, D95 and S97 are located in a disordered section in this model but would close in the actual active site in the actual structure. It has been proposed that Asp101 (95 in the AlphaFold structure) deprotonates Arg220 (214 in the model), which then acts as a nucleophile which attacks the phosphate group of phosphatidylcholine. Given the very high pKa of arginine, this mechanism, if true, is somewhat unique. Phosphocholine is linked to ceramide to produce sphingomyelin. Three key and extremely conserved amino acids in the active site are Asp101, Arg220 and Asn358. A specific ceramide transport molecule (CERT) is important to the reaction with SMS1 in that it transfers ceramide from the cytosolic surface of the endoplasmic reticulum to the trans-Golgi in an ATP-dependent and non-vesicular manner. Much of the sphingomyelin produced in the Golgi is then delivered to the apical plasma membrane by a vesicular transport mechanism. Sphingomyelin synthesis is regulated in part by phosphatidylinositide metabolism and is connected to sterol homeostasis through the oxysterol binding protein (OSBP). SMS2 in the plasma membrane is not dependent on CERT-mediated ceramide delivery, but is believed to convert ceramide produced locally by a sphingomyelinase back to sphingomyelin; this may be an important protective mechanism for the cell. The location of the enzymes explains the enrichment of sphingomyelin in specific membranes and the sidedness, i.e., the luminal trans-Golgi and the outer leaflet of the plasma membrane, while ceramide reaching the cis-Golgi is utilized for the synthesis of glucosylceramide. As the nature of the molecular species of sphingomyelins produced differs appreciably from that of the ceramide precursors, the sphingomyelin synthases must have considerable substrate specificity. The reaction can be reversible, using sphingomyelin to generate ceramide for specific signaling functions. It is evident that sphingomyelin biosynthesis forms a link between the sphingolipid signaling pathway (pro-apoptotic - see below) and that of glycerolipids via the mitogenic diacylglycerol by‑products. Although the importance of this production relative to that via phosphatidylinositol is not known, it is possible that it is significant locally at the external leaflet of the plasma membrane. An alternative pathway of sphingomyelin synthesis has been demonstrated in the endoplasmic reticulum in which ceramide is first converted to ceramide phosphoethanolamine via transfer of the head group from phosphatidylethanolamine, followed by stepwise methylation of the ethanolamine moiety. However, the physiological significance of this pathway has yet to be established. It was long thought that the only function of sphingomyelin was to serve as a substitute for phosphatidylcholine as a building block of membranes, i.e., by forming a stable and chemically resistant outer leaflet of the plasma membrane lipid bilayer. For example, it may limit the ingress of oxygen and thence oxidation of adjacent unsaturated acyl chains. While this is certainly one of its functions, the apparent similarity between phosphatidylcholine and sphingomyelin is superficial, and there are great differences in the hydrogen bonding capacities and physical properties of the two lipids. For example, sphingomyelin has an amide bond at position 2 and a hydroxyl at position 3 of the sphingoid base, both of which can participate in hydrogen bonding, while the trans double bond also appears to assist intermolecular interactions in membranes. Indeed, the first five carbon atoms of the sphingoid base in sphingolipids constitute a key feature that has been termed the ‘sphingoid motif’, which facilitates a relatively large number of molecular interactions with other membrane lipids, via hydrogen-bonding, charge-pairing, hydrophobic and van der Waals forces. With phosphatidylcholine, in contrast, the two ester carbonyl groups can act only as hydrogen acceptors. The degree of unsaturation of the alkyl moieties in each lipid is very different, and this gives them dissimilar packing properties in membranes. It is now recognized that sphingomyelin and other sphingolipids have a strong tendency to interact with proteins and cholesterol, often via strong van der Waals interactions and hydrogen bonding, to form transient nano-domains in membranes known as 'rafts' and on the surface of lipoprotein particles. Initially, there was a view that saturated sphingomyelin formed a liquid-ordered phase with cholesterol or a gel phase with saturated ceramides to lead to lateral segregation within the membrane, and that sphingomyelin and cholesterol metabolism were closely integrated, even that the sphingomyelin concentration might control the distribution of cholesterol in cells. On the other hand, the understanding of the mechanism of raft formation in membranes has changed substantially in recent years, and while an interaction with cholesterol is certainly important, it may not be the major factor in vivo. Ceramide can displace cholesterol from its association with sphingomyelin, when formed in membranes by hydrolysis of the latter. Other functions: Sphingomyelin per se is generally considered to be a relatively inert molecule, although modern molecular biology methods are uncovering potential regulatory functions via interactions with particular proteins. For example, it has been shown to inhibit the activity of phospholipase A2α, a key enzyme in eicosanoid production. Sphingomyelin in the plasma membrane may be essential for the internalization of transferrin and thence of iron into cells, and it appears to be required for the activity of a number of membrane-bound proteins, including those of certain ion channels and receptors. As the most abundant sphingolipid in the nucleus, it is intimately involved in chromatin assembly and dynamics as well as being an integral component of the nuclear matrix. A single molecular species of sphingomyelin with a C18 acyl chain binds specifically to a coat protein designated 'p24' to enable it to form membrane vesicles. In addition, sphingomyelin is selectively recognized and acts as a receptor for the actinoporins, which are pore-forming toxins produced by sea anemones. There is a specific binding site for sphingomyelin on the amyloid beta-peptide (Aβ) in brain, and there is evidence from studies in vitro that this may promote the aggregation of these proteins in Alzheimer's disease. In turn, this leads to depletion of brain sphingomyelin by activation of acid sphingomyelinase with disruption of many protein–lipid interactions and thence of downstream signaling pathways. In contrast, the ganglioside GM1 may have a protective role towards Aβ aggregation As well as its role in membranes, it serves as a precursor for ceramides, long-chain bases, sphingosine-1-phosphate, and many other biologically important sphingolipids, as part of the 'sphingomyelin cycle' (also termed the ‘sphingolipid’ or ‘ceramide’ cycles depending on the context). Some of these metabolites are intra- and inter-cellular messengers, and others are essential membrane constituents. The sphingomyelin cycle extends to other sphingolipids via the action of sphingomyelinases and enzymes such as glycosylhydrolases and glycosyltransferases in cells to produce innumerable new oligoglycosylceramides. It can also give rise to sn-1,2-diacylglycerols, which are central to many metabolic and signaling pathways. These molecular relationships are illustrated only briefly in Figure \(23\). In particular, sphingomyelin is a major source of ceramides in most cellular organelles, including the nucleus and even mitochondria, via the action of sphingomyelinases (see next section), and in addition to being a source of other sphingolipids these are required to trigger apoptosis and other metabolic changes. As ceramides do not mix well with glycerophospholipids and cholesterol, this conversion results in the formation of new membrane domains enriched in ceramide that exclude cholesterol and so differ in composition from other sphingolipid rafts. This has profound effects on membrane function, especially of the plasma membrane, in that different proteins may be recruited or excluded depending on their relative affinities for cholesterol and ceramides. It may also influence disease states such as cancer. Chlamydiae, widespread bacterial pathogens, acquire sphingomyelin from the Golgi apparatus and plasma membrane of their hosts and this is necessary for the viability and growth of the organisms. Other pathogenic bacteria, notably Pseudomonas aeruginosa and Neisseria gonorrhoeae, can hijack sphingomyelin catabolic enzymes with deleterious effects upon the host. Likewise, human immunodeficiency virus (HIV) and the hepatitis C virus utilize host sphingomyelin for their own nefarious purposes. Nutrition: Although there is no known nutritional requirement for sphingomyelin and other sphingolipids, they are a component of any diet containing egg, meat or dairy products. Thus, it has been estimated that per capita sphingolipid consumption in the United States, for example, is of the order of 0.3-0.4 g/d. As sphingolipids constitute an appreciable proportion of the polar lipid constituents of milk, they may be significant if minor nutrients for infants and beneficial effects upon their development have been claimed. From animal experiments, there is evidence that dietary sphingolipids can reduce the intestinal absorption of cholesterol and other lipids, leading to reductions in serum lipid concentrations. Feeding sphingolipids inhibits colon carcinogenesis and may alleviate some of the symptoms of inflammatory bowel disease. 2-Hydroxyoleic acid suppresses the growth and induces autophagy in cancer cells by stimulating the synthesis of sphingomyelin and increasing the amount of this lipid in the plasma membrane. On the other hand, plasma sphingomyelin levels are considered to be an independent risk factor for atherosclerosis, possibly as a result of its ability to retain cholesterol in cells and the arterial wall with consequent diminished reverse cholesterol transfer via HDL. Sphingomyelin Catabolism In contrast to the glycerolipids, dietary sphingolipids are not hydrolyzed by pancreatic enzymes only. Rather, most of the sphingomyelin in the diet is hydrolyzed in the brush border of the intestines by an alkaline sphingomyelinase (at a pH of 8.5–9 optimally) to ceramide and thence by a neutral ceramidase to free fatty acids and sphingosine. Some of this enzyme is also present in liver from which it is secreted in bile into the intestinal lumen where it can hydrolyze sphingomyelin and other phospholipids with the aid of bile salts. The sphingosine released at the brush border is absorbed, some is re-N-acylated to form ceramides, and the remainder is converted via sphingosine-1-phosphate to palmitic acid, which is esterified into the triacylglycerol component of chylomicrons. In the process, some of these sphingolipid intermediates may have signaling functions and anti-inflammatory properties in intestinal cells. The alkaline sphingomyelinase is unusual in that is very different in its structure and other properties from intracellular enzymes with a related function; it is part of the (ecto)nucleotidepyrophosphatase-phosphodiesterase protein family (NPP) that includes autotaxin. The enzyme is believed to have a role in the production of sphingolipid metabolites within the intestines and colon especially, which may influence a number of disease states. For example, it appears to inhibit colon cancer by generating ceramides. In addition, alkaline sphingomyelinase has phospholipase C activity towards the pro-inflammatory metabolite platelet-activating factor and towards lysophosphatidylcholine with potentially further beneficial effects. By reducing the level of endogenous sphingomyelin and increasing that of ceramides in the membranes of intestinal cells, it is believed to reduce the uptake of dietary cholesterol. Catabolism in other tissues: The key enzymes for the degradation of sphingomyelin to ceramides in most tissues are also sphingomyelinases (phosphodiesterases), as shown in Figure \(24\). These are similar in function to phospholipase C and generate ceramides with their innumerable and important signaling properties as the main product. There are many such enzymes with different pH optima and metal ion requirements that operate in different regions of the cell with potentially distinct biochemical roles. Thus, there is an acid sphingomyelinase in the endo-lysosomes, and different neutral sphingomyelinases in the plasma membrane, endoplasmic reticulum, Golgi, and mitochondria in addition to the alkaline sphingomyelinase in the intestines. It should not be forgotten that the other product of the reaction is phosphocholine, which has importance as a nutrient. Bacterial sphingomyelinases are known to lyse red blood cells, although intriguingly, there is a sphingomyelinase in the bacterium Pseudomonas aeruginosa that can also act as a sphingomyelin synthase in vitro at least. The lysosomal acid sphingomyelinase (pH optimum ca. 5) is expressed ubiquitously and has a key housekeeping role in maintaining normal membrane turnover and remodeling of the sphingolipid constituents, especially those of lipoproteins. While other lysosomal sphingolipid hydrolases require a saposin activator protein for full activity, the acid sphingomyelinase incorporates a built-in N-terminal saposin domain so does not require an external activator. Under resting conditions, acid sphingomyelinase is stored inside lysosomes, but upon stimulation, it undergoes vesicular transport to the plasma membrane where it docks with a specific protein and is exposed on the outer leaflet. It then generates ceramide by hydrolysis of sphingomyelin and initiates the train of events that leads to apoptosis. There are reports that acid sphingomyelinase, by acting at the plasma membrane to produce ceramides, regulates the localization and trafficking of palmitoylated proteins from the Golgi, and it may also facilitate bacteria-host interactions. Experiments in vitro have demonstrated that the enzyme can be considered as a phospholipase C that is active against a wide range of phospholipids, including ceramide-1-phosphate and the unique lysosomal phospholipid bis(monoacylglycero)phosphate. There is a related secreted acid sphingomyelinase (Zn2+-dependent), which can be transported to the outer membrane of the cell and is especially important in endothelial cells of the human coronary artery. This enzyme is produced by the same gene but differs from the lysosomal enzyme as it requires Zn2+ ions for activation and has a different glycosylation pattern. It can also operate at neutral pH and has multiple functions in that it is involved in many aspects of cellular signaling as well as in membrane sphingomyelin turnover. By acting at the plasma membrane to produce ceramides, it is believed to regulate the trafficking of palmitoylated proteins from the Golgi to their new location. Neutral sphingomyelinases (pH optima 7.4), of which four quite distinct enzymes are known, are located in membranes of the endoplasmic reticulum, Golgi, and plasma membrane with one in mitochondria (MA-NSM), where they have signaling functions by generating ceramides and thence other biologically active sphingolipids. Human NSM-1 has 423 amino acid residues and a molecular weight of 47.6 kDa; it has two putative transmembrane domains in the C-terminus and resides mainly in the nucleus and endoplasmic reticulum. It has a broad specificity for choline phospholipids, but it is most active with sphingomyelin and may not have a significant role in cellular signaling. In contrast, NSM-2 which is located in the Golgi apparatus and plasma membrane is activated by phosphatidylserine and is important for ceramide signaling. It is especially important in brain and nervous tissue, where it is required for the secretion of hypothalamic-sssreleasing hormones, although it is relevant to many cellular functions and physiological processes in most other tissues. Dysregulation of NMS-2 is reported to be a factor in many inflammation-related pathologies. Neutral sphingomyelinases-3 is found mainly in the plasma membrane of bone and cartilage, where it is vital for the process of mineralization; it is also important in striated and cardiac muscle. Little seems to be known of the function of the mitochondrial enzyme. Losses, mutation, and poor expression of the gene encoding neutral sphingomyelinase have been observed in several cancers, but exposure to ionizing irradiation led to rapid hydrolysis of sphingomyelin to ceramide by this enzyme, and thence to cancer cell death. A diverse range of factors activates the enzymes, including chemotherapeutic agents, tumor necrosis factor-alpha, 1,25-dihydroxy-vitamin D3, endotoxin, gamma-interferon, interleukins, nerve growth factor, and most conditions known to induce cellular stress, especially in relation to inflammation. As they utilize by far the most abundant sphingolipid in animal tissues to generate ceramides and other sphingolipid metabolites that have important signaling functions, sphingomyelinases are believed to function as regulators of signaling mechanisms, especially in the nucleus of the cell. Thus, they have a much wider metabolic role than simply catabolism of sphingomyelin. The type A and B forms of Niemann-Pick disease are lysosomal lipid storage disorders that are a consequence of a deficiency of acid sphingomyelinase with a resulting accumulation of sphingomyelin and smaller amounts of other sphingolipids, including gangliosides, in cells and tissues and especially in the monocyte/macrophage system to form the so-called “foam cells” that characterize the disease. A consequent lack of ceramide production may be involved in the pathology of the disease. Increasing sphingomyelin levels in turn result in elevated cholesterol concentrations. It is noteworthy that membranes containing ceramides have a much lower binding capacity for cholesterol, so sphingomyelin degradation may play a part in cholesterol homeostasis. Type C Niemann-Pick disease differs from the A and B forms and is caused by defects in two distinct cholesterol-binding proteins (NPC1 and NPC2). Glucosyl- and Galactosylceramides (Cerebrosides) There are two natural monoglycosylceramides of special importance in animals, i.e., glucosylceramide and galactosylceramide. Both have biological functions in their own right, but especially as structural components of membranes, as in the brain, for example, where galactosylceramide is required for the maintenance of the structure and stability of myelin and the differentiation of oligodendrocytes. Glucosylceramide is a vital component of all cell types, and is most abundant in human skin; it is the key intermediate in the biosynthesis of lactosylceramide and thence of complex oligoglycosphingolipids, including gangliosides. This monoglycosylceramide is also a major component of the membranes of plants and fungi. Although the two lipids have very similar structures in that D-galactose is an epimer of D-glucose and they differ only in the configuration at C4, they have very different biological properties. A few other monoglycosylceramides are produced in nature, for example by some bacteria of the order Sphingomonadales of α‑proteobacteria. Structure and Occurrence β-D-Galactosylceramide (Galβ1-1'Cer) is the principal glycosphingolipid in brain tissue, hence the trivial name "cerebroside", which was first conferred on it in 1874, although it was much later before it was properly characterized. In fact, galactosylceramides are found in all nervous tissues and indeed at low levels in all organs, but in they brain they can amount to 2% of the dry weight of grey matter and 12% of white matter or 23% of myelin lipids, where they insulate the axons of neuronal cells and constitute a substantial component of the extended plasma membrane of oligodendrocytes. It is also present in some fungal species. While galactosylceramide can be sulfated to form a sulfatide or sialylated to form ganglioside GM4, only a small proportion is subjected to further galactosylation to form Gal2Cer as the precursor for the limited gala-series of oligoglycosphingolipids. β-D-Glucosylceramide (Glcβ1-1'Cer), with the trivial name "glucocerebroside", is a major constituent of skin lipids, where it is essential for the maintenance of the water permeability barrier of the skin. Otherwise, it is most abundant in animal tissues such as the spleen and erythrocytes as well as in nervous tissues, especially in the neurons if at low levels, and it is also found in plants. Higher than normal concentrations of this glycosphingolipid have been reported for the apical plasma membrane domain of epithelial cells from the intestines (especially the absorptive villous cells) and urinary bladder. The d18:1/16:0 molecular species of the two lipids are illustrated in Figure \(25\). However, of equal or greater importance to the natural occurrence of glucosylceramide per se is its role as the biosynthetic precursor of lactosylceramide in animals, and thence of most of the complex neutral oligoglycolipids and gangliosides. In contrast, glucosylceramide is the end-product of the biosynthetic pathway in plants and fungi. Interestingly, the proportion of galactosylceramides relative to glucosylceramides in myelin glycolipids increases greatly in the ascending phylogenic tree, and the ratio of hydroxy- to nonhydroxy fatty acids in cerebrosides increases with the complexity of the central nervous system. There is also an intriguing sex difference in the kidney, where it has been shown that galactosylceramide rather than glucosylceramide occurs in male mice only (or androgen-treated adult females). Only glucosylceramide is present in the nerves of the most primitive animals (protostomes). In the brain, the galactosylceramides are enriched in very-long-chain fatty acids (C22–C26). The fatty acid and long-chain base compositions of cerebrosides from the intestines of the Japanese quail are listed in Table \(5\) for illustrative purposes. The fatty acid components resemble those of other sphingolipids, although the percentage of 2-hydroxy acids is higher than that in sphingomyelin, for example. They are exclusively saturated in this instance, though a small proportion of monoenoic components may also be found in other tissues. Glucosylceramides tend to contain mainly non-hydroxylated fatty acids that are of relatively shorter chain length. The proportion of trihydroxy bases listed is perhaps higher than in other many other tissues or species studied, probably reflecting the diet. Usually, sphingosine is the main long-chain base in cerebrosides of animal tissues. Table \(5\): Composition of fatty acids and long-chain bases (wt % of the total) in cerebrosides of intestines from the Japanese quail.* Long‑chain bases Fatty acids Non-hydroxy acids 2-Hydroxy acids Type %   % % t18:0 43 16:0 5 6 d18:0 9 18:0 3 trace d18:1 27 20:0 2 4 t20:0 6 21:0 trace 2 d20:0 3 22:0 4 43 d20:1 11 23:0 1 13 24:0 3 12 * The cerebrosides comprised 81% galactosylceramide and 19% glucosylceramide. From Hirabayashi, Y. et al., Lipids, 21, 710-714 (1986); DOI. Plants: Glucosylceramide is the only glycosphingolipid common to plants, fungi, and animals. It has often been described incorrectly as the main sphingolipid in plants, but this has been because the more polar complex glycosylinositol phosphoceramides are not easily extracted and until relatively recently were missed in conventional analyses. Nonetheless, glucosylceramide is abundant in photosynthetic tissues and constitutes approximately a third of the total sphingolipids, where the main long-chain bases are C18 4,8‑diunsaturated (Z/Z and E/Z) (not sphingosine as illustrated above); it is a major component of the outer layer of the plasma membrane and is also enriched in the late endosomes and plant tonoplast. Small amounts of monoglycosylceramides containing a β‑D‑mannopyranosyl unit may be present in non-photosynthetic tissues, but galactosylceramides have not been found in plants. Glucosylceramide is a common component of the lipids of yeast and other fungi, including most fungal pathogens. However, it does not occur in the yeast Saccharomyces cerevisiae, which is widely used as an experimental model, although trace levels of galactosylceramide have been detected. The fatty acid and long-chain base compositions of glucosylceramides from two plant sources are listed in Table \(6\). Perhaps surprisingly, the fatty acid components are not very different in nature from those in animal tissues, comprising mainly longer-chain saturated and monoenoic acids, with a high proportion being saturated and having a hydroxyl group in position 2. In the examples selected for the table here, both di- and tri-hydroxy long-chain bases were found, mainly diunsaturated (Z/Z and E/Z) and almost entirely C18 in chain length. Much higher concentrations of glucosylceramides are found in pollen than in leaves, with substantial compositional differences. For example, the long-chain bases in Arabidopsis leaves consist mainly of t18:1, with relatively little d18:1, t18:0 and d18:0 (with 16:0, 24:0 and 24:1 hydroxy fatty acids mainly), but no d18:2 base although this is 50% of those in pollen. While saturated 2-hydroxy acids predominate in most plants, some cereal glucosylceramides contain high proportions of mono-unsaturated very-long-chain fatty acids of the n-9 family. Glucosylceramides from algae tend to resemble those from higher plants, although some novel structures have been reported from microalgae. Table \(6\): Composition of fatty acids and long-chain bases (wt % of the total) in glucosylceramides of seeds from scarlet runner beans and kidney beans. Fatty acidsa Long-chain basesb Type Runner beans Kidney beans   Runner beans Kidney beans % %   % % 16:0 4 5 t18:0 trace trace Other non-hydroxy 1 2 t18:1-8t 13 11 14:0-OH 1 1 t18:1-8c 10 9 15:0-OH 1 1 d18:0 trace trace 16:0-OH 58 58 d18:1-8c/t 1 3 18:0-OH trace trace d18:1-4t trace trace 20:0-OH trace trace d18:2-4t,8t 45 60 22:0-OH 7 6 d18:2-4t,8c 31 17 23:0-OH 2 1 24:0-OH 23 23 25:0-OH 1 1 26:0-OH 1 1 From Kojima, M. et al., J. Agric. Food. Chem., 39, 1709-1714 (1991); DOI, but see also Yamashita, S. et al. for further data: DOI a including 2-hydroxy acids; b di- and tri-hydroxy bases with cis or trans double bonds in the positions indicated. Biosynthesis Ceramides synthesized both de novo and by catabolism of sphingomyelin are used for the biosynthesis of monoglycosylceramides in animal tissues. The biosynthetic mechanism resembles that for glycosyldiacylglycerols, i.e., there is a direct transfer of the carbohydrate moiety from a sugar-nucleotide, e.g. uridine 5-diphosphate(UDP)-galactose, UDP-glucose, etc, to a ceramide unit synthesized in the endoplasmic reticulum. This is illustrated in Figure \(26\). During the transfer, which is catalyzed by specific glycosyl-transferases, inversion of the glycosidic bond occurs from the alpha to beta configuration. Synthesis of β-D-galactosylceramide takes place on the lumenal surface of the endoplasmic reticulum, although it has free access to the cytosolic surface by an energy-independent flip-flop process. Expression of the UDP-galactose:ceramide galactosyl transferase (galactosylceramide synthase) is restricted to oligodendrocytes, Schwann cells, kidneys, and testes. Prior to sulfation, galactosylceramide is transported to the trans-Golgi compartment. In contrast, after the transfer of the precursor ceramides from the endoplasmic reticulum to the cytosolic side of the early Golgi membranes with the aid of the CERT protein, glucosylceramide is produced by a glucosylceramide synthase present in this membrane (with the possible exception of neuronal tissues). If it is to be converted to more complex oligoglycosylceramides, this must be translocated to the luminal leaflet of the trans-Golgi membranes, a process that occurs both by vesicular and by non-vesicular transport. The latter is mediated by a conserved clade of integral membrane proteins, i.e., phospholipid flippases (P4-ATPases) designated ATP10A and ATP10D, together with the four phosphate adapter protein-2 (FAPP2) and glycolipid transfer protein (GLTP) in humans with related enzymes in fungi, which utilize the energy from ATP catalysis to translocate lipids across cellular membranes. The human enzymes are entirely specific for glucosylceramide and not galactosylceramide. Indeed, the galactosyl- and glycosylceramide synthases have no significant sequence homology, indicating different evolutionary origins. For their functions in protein interactions and signaling, both galactosyl- and glucosylceramide must be transported to and then across the plasma membrane. Some glucosylceramide is carried by lipoproteins (VLDL, LDL, and HDL) in the circulation and presumably requires active transport for absorption and distribution across the membranes of target tissues. In plants, glucosylceramides are also formed by an evolutionarily conserved glucosylceramide synthase involving UDP-glucose in the endoplasmic reticulum, although an alternative mechanism has been described that utilizes sterol glucoside as the immediate glucose donor to ceramide. There is also evidence for a requirement for ceramides containing Δ4 trans-double bonds for synthesis of glucosylceramides but not other sphingolipids in some plant and fungal tissues. However, there is a distinct ceramide synthase in the yeast Pichia pastoris, which produces ceramides of defined composition exclusively for the production of glucosylceramides. A separate ceramide synthase with different specificities produces the ceramide precursors for ceramide phosphorylinositol, which contains only phytosphingosine as the long-chain base. In fungi, glucosylceramide synthases have been characterized, but a galactosylceramide synthase has yet to be identified. Enzymes responsible for the biosynthesis of glucuronosylceramide and α-galactosylceramide in some bacterial species have been characterized. Function Galactosylceramides: A remarkable property of cerebrosides is that their 'melting point' is well above physiological body temperature, so that glycolipids have a para-crystalline structure at this temperature. Each cerebroside molecule may form up to eight inter- or intramolecular hydrogen bonds by lateral interaction between the polar hydrogens of the sugar and the hydroxy and amide groups of the sphingosine base of the ceramide moiety, and this dense network of hydrogen bonds is believed to contribute to the high transition temperature and the compact alignment of cerebrosides in membranes. As with sphingomyelin, monoglycosylceramides tend to be concentrated in the outer leaflet of the plasma membrane together with cholesterol and thence in myelin in the specific membrane domains termed 'rafts'. Indeed, the latter appear to facilitate segregation to a greater extent than sphingomyelin via the combination of hydrogen bonds and hydrophobic interactions, and these forces are also of great importance for binding to the wide range of proteins, including enzymes and receptors, which are found in raft domains. Galactosylceramide is essential to myelin structure and function and it is involved in oligodendrocytes differentiation. While molecular species with 2’-hydroxy fatty acid constituents are not essential for myelin formation, they are critical for the long-term stability of myelin, presumably because increased hydrogen bonding with neighboring lipids in membranes stabilizes the phase structure. Galactosylceramide is important as a precursor of 3’-sulfo-galactosylceramide, which is also essential to brain development in addition to numerous functions in other tissues. By interacting with sulfatide located in the membrane of opposing layers in the myelin sheath by carbohydrate-carbohydrate interaction, it forms what is known as a glycosynapse, which provides a necessary contribution to the long-term stability of myelin. Glucosylceramides: Glucosylceramides have similar physical properties in membranes to the galactose analog, and they are also concentrated in raft domains in the outer leaflet of the plasma membrane. As mentioned briefly above, they are the primary precursor for most of the more complex oligoglycosphingolipids in animal tissues, especially in brain, where synthesis is vital for the production of most neuronal oligoglycosphingolipids, while glucosylceramide per se is essential for axonal growth. They are major constituents of skin lipids, where they are essential for lamellar body formation in the stratum corneum and to maintain the water permeability barrier of the skin. In addition, the epidermal glucosylceramides (together with sphingomyelin) are the source of the unusual complex ceramides that are found in the stratum corneum including those with terminal hydroxyl groups and estolide-linked fatty acids. Some of the glucosylceramide in the skin is linked covalently to proteins via terminal hydroxyl groups, presumably to strengthen the epidermal barrier. Much of the evidence for the function of glycosylceramides in animals has been derived from cell lines in which synthesis of the lipid has been suppressed by various means in vitro. It appears that glucosylceramide is not essential for the viability of certain cell lines in culture, but disruption of the global synthase gene in mice results in the death of embryos. It is essential for the survival of cancer cells, and deletion from other cell types can lead to abnormalities. In addition to being an intermediate in the biosynthesis of more complex glycosphingolipids and its role in the permeability barrier of the skin (discussed above), glucosylceramide is believed to be required for intracellular membrane transport, cell proliferation, and survival, and for various functions in the immune system. In contrast, there are indications that it may have adverse implications for various disease states. For example, over-expression of glucosylceramide synthase in cancer cells has been linked to tumour progression with a reduction in ceramide concentration, resulting in increased resistance to chemotherapy. The lipid has also been associated with drug resistance in a wider context. In the nematode Caenorhabditis elegans, glucosylceramide containing the fatty acid 22:0 is reported to be a longevity metabolite that functions through the membrane localization of clathrin, a protein that regulates membrane budding. In Arabidopsis, glucosylceramides are critical for cell differentiation and organogenesis, but not necessarily for the viability of cells. It has been proposed that glycosphingolipids could impose positive curvature to membranes, thereby facilitating vesicle fusion. There is evidence that glycosylceramides (but not glycosyldiacylglycerols) together with sterols are located in 'rafts' in plant membranes in an analogous manner to sphingolipids in animal tissues, and that they are associated with specific proteins. Correlative studies suggest that glucosylceramides help the plasma membrane in plants to withstand stresses brought about by cold and drought. For example, glycosylceramides containing 2-hydroxy monounsaturated very-long-chain fatty acids and long-chain bases with 4-cis double bonds appear to be present in higher concentrations in plants that are more tolerant of chilling and freezing. While fungal glucosylceramides with a 9-methyl group within the sphingosine backbone elicit defence responses in rice, cerebrosides with double bonds in positions 4 and/or 8 of the long-chain base appear to be involved in the defense of some plant species against fungal attack. Less is known of the function of glucosylceramides in fungi, although they are certainly major constituents of the plasma membrane and cell wall. They are believed to be involved in such processes as cell wall assembly, cell division and differentiation, and signaling. The presence of the methyl branch in the long-chain base is essential for cell division and alkali tolerance. In the case of fungal pathogens, glucosylceramides are recognized by the host immune system and regulate virulence, often after export into the external environment as extracellular vesicles. In contrast to animals, ceramide monohexosides are not precursors for oligoglycosylceramides in fungi. Some molecular species of this lipid from plants (a Δ8 double bond in the long-chain base is essential) show fruiting-inducing activity in the fungus Schizophyllum commune. α-D-Galactosylceramides: Cerebrosides linked to an α-D- rather than a β-D-galactosyl unit such as that found in the marine sponge Agelas mauritianus, in human gut microflora, and even in cow's milk are potent stimulators of mammalian immune systems by binding to the protein CD1d on the surface of antigen-presenting cells and activating invariant natural killer T cells. Indeed this was one of the first pieces of evidence to show that glycolipids, like glycoproteins, could invoke an immune response. Subsequently, it was demonstrated that α-galactosylceramide with a 24:1 fatty acid, though present in very small amounts, is loaded onto the CD1d or CD40 protein and is presented as the natural endogenous ligand for NKT cells in the thymus and the periphery. Once activated, NKT cells secrete a range of pro-and anti-inflammatory cytokines to modulate innate and adaptive immune responses. The α‑glucosyl and α‑psychosine analogs show similar activity. It is not certain whether α-galactosylceramide is synthesized in animal tissues, and it is likely that is derived primarily from members of the gut microbiome such as Bacteroides fragilis and related species (although in general, few bacterial species produce sphingolipids). Ceramide-galactosyltransferases responsible for the synthesis of this lipid in two species of bacteria from the intestinal microbiome have been identified. In mouse gut, the main molecular form consisted of a 2‑(R)‑hydroxylated hexadecanoyl chain linked to C18-sphinganine, while that in B. fragilis contained longer-chain components with iso-methyl-branches in the sphingoid base and often fatty acid moieties. The sphinganine chain branching is a critical determinant of NKT cell activation by the bacterial enzyme. A decrease in the production of this lipid was observed in mice exposed to stress conditions that alter the composition of the gut microbiota, including Western-type diet, colitis, and influenza A virus infection with potential consequences upon the systemic immune responses. Its concentration within animal tissues is controlled by catabolic enzymes in a two-step mechanism: removal of the acyl chain by an acid ceramidase followed by hydrolysis of the sugar residue by an α-glycosidase. Initial studies with animal models suggest that treatment with α‑D‑galactosylceramides is effective against lung and colorectal cancers, melanomas and leukemia, and pre-clinical trials of this lipid and synthetic analogs so far have shown that these are safe and effective as an anti-tumour immunotherapeutic agents and vaccine adjuvants. Indeed, a phase I trial with high-risk melanoma patients has given promising preliminary results. Catabolism of Glycosphingolipids In animal tissues, the main sites for the degradation of all glycosphingolipids, including the monoglycosylceramides, oligoglycosphingolipids and gangliosides, are the lysosomes. These are membrane-bound organelles that comprise a limiting external membrane and internal lysosomal vesicles, which contain soluble digestive enzymes that are active at the acidic pH of this organelle. All membrane components are actively transported to the lysosomes to be broken down into their various primary components. In the case of glycosphingolipids, this means to fatty acids, sphingoid bases, and monosaccharides, which can be recovered for re-use or further degraded. Thus, sections of the plasma membrane enter the cell by a process of endocytosis, and they are then transported through the endosomal compartment to the lysosomes. The compositional and physical arrangement of the lysosomal membranes is such that they are themselves resistant to digestion with bis(monoacylglycero)phosphate (lysobisphosphatidic acid) as a characteristic component of the inner membrane. A glycocalyx of highly N-glycosylated integral membrane proteins protects the perimeter membrane with the aid of the ganglioside GM3, which is resistant to degradation. This glycocalix forms an efficient hydrophilic barrier at the luminal surface of the lysosomal perimeter membrane to protect it from degradation by proteases and hydrolases, and to prevent lipids and their hydrolysis products from escaping from the lumen of the lysosome. Degradation of oligoglycosylceramides and gangliosides occurs by sequential removal of monosaccharide units via the action of specific exohydrolases from the non-reducing end until a monoglycosylceramide unit is reached when glucosylceramide β-glucosidases or an analogous β-galactosidase (one isoform) removes the final carbohydrate moiety. Several glucosylceramidases are known; GBA1 is a lysosomal hydrolase, GBA2 is a ubiquitous non-lysosomal enzyme and GBA3 is a cytosolic β-glucosidase. The last is found in the kidney, liver, spleen and a few other tissues of mammals, but its function is not clear. As glycolipids with fewer than four carbohydrate residues are embedded in intralysosomal membranes, while the degradative enzymes are soluble, the process requires the presence of negatively charged lipids and specific activator proteins, which are water-soluble glycoproteins of low molecular weight. These are not themselves active catalytically but are required as cofactors either by directing the enzyme to the substrate or by activating the enzyme by binding to it in some manner. Five such proteins are known, the GM2-activator protein (specific for gangliosides) and Sphingolipid Activator Proteins or saposins A, B, C and D, which perturb the membranes sufficiently to enable the degradative enzymes to reach the glycolipid substrates. The four saposins are derived by proteolytic processing from a single precursor protein, prosaposin, which is synthesized in the endoplasmic reticulum, transported to the Golgi for glycosylation, and then to the lysosomes. Of these, saposin C is essential for the degradation of galactosyl- and glucosylceramide, while saposin B is required for the hydrolysis of sulfatide, globotriaosylceramide, and digalactosylceramide. The products of the hydrolysis reaction with monoglycosylceramides are ceramides and monosaccharides with net retention of the stereochemistry of the latter in the process. This is illustrated in Figure \(26\). The reactions are aided by the presence of anionic lipids such as bis(monoacylglycero)phosphate. In particular, this increases the ability of the GM2-activator to solubilize lipids and stimulates the hydrolysis of membrane-bound GM1, GM2, and some of the kidney sulfatides. Saposin D stimulatesthe  degradation of lysosomal ceramide by acid ceramidase, and it is also involved in the solubilization of negatively charged lipids at an appropriate pH. Eventually, the ceramides can in turn be hydrolyzed by an acid ceramidase to fatty acids and sphingoid bases. β-Glucosylceramidase and saposin C are also required for the generation of the structural ceramides from glucosylceramide in the outer region of the skin, a process essential for optimal skin barrier function and survival. Some glucosylceramide is hydrolyzed by the enzyme GBA2 at the plasma membrane, where the ceramide formed is rapidly converted to sphingomyelin by the sphingomyelinase 2, which may be co-located with the glucosidase. In addition, it has been established that cellular β-glucosidases are able to transfer the glucose moiety from glucosylceramide to and from other lipids as in the formation of cholesterol glucoside. Small but significant amounts of glucosyl- and galactosylceramides are ingested as part of the human diet. They are not hydrolyzed by pancreatic enzymes but are degraded in the brush border of the intestines by the enzyme lactase-phlorizin hydrolase (which also hydrolyses the lactose in milk) to ceramides and thence to sphingosine. An Arabidopsis homolog of human glucosylceramidase (AtGCD3) preferentially hydrolyses glucosylceramides that contain long acyl chains, and three further isoforms may exist based on sequence homology. Genetic disorders and Disease Harmful quantities of glucosylceramide accumulate in the spleen, liver, lungs, bone marrow, and, in rare cases, the brain of patients with Gaucher disease, the most common of the inherited metabolic disorders (autosomal recessive) involving storage of excessive amounts of complex sphingolipids. Three clinical forms (phenotypes) of the disease are commonly recognized of which by far the most dangerous are those affecting the brain (Types 2 and 3). All of the patients exhibit a deficiency in the activity of the lysosomal glucosylceramide-β-glucosidase (GBA1), which catalyzes the first step in the catabolism of glucosylceramide. The enzyme may be present, but a mutation prevents it from forming its correct conformation, although other factors may be involved as patients with a defective saposin C, the lysosomal activator protein, develop similar symptoms. In the brain, glucosylceramide accumulates when complex lipids turn over during brain development and during the formation of the myelin sheath of nerves. Other than in the brain, the excess glucosylceramide arises mainly from the biodegradation of old red and white blood cells. The result is that the glucosylceramide remains stored within the lysosomes of macrophages, i.e., the specialized cells that remove worn-out cells by degrading them to simple molecules for recycling, thus preventing them from functioning normally and often leading to chronic inflammation. The enlarged macrophages containing undigested glucosylceramide are termed Gaucher cells. They over-express and secrete certain proteins into the circulation, and some of these are used as biomarkers. In addition, glucosylceramide is converted more rapidly to gangliosides in these cells, leading to an increase in ganglioside GM3 in the plasma and spleen of patients with Gaucher disease. Fortunately, there are now effective enzyme replacement therapies for patients with the milder (non-neurological or Type 1) form of Gaucher disease that successfully reverse most manifestations of the disorder, including decreasing liver and spleen size and reducing skeletal abnormalities. Two oral drugs that inhibit glucosylceramide synthesis have also been approved. Defective GBA1 enzyme activity in humans has been implicated in an increased risk of multiple myeloma and other cancers. Oligoglycosylceramides and gangliosides in particular are known to be involved in the pathology of a number of cancers, and glucosylceramide is an important precursor of these. Inhibition of glucosylceramide synthase, which is overexpressed in many human tumors lead to a marked arrest of cell growth in cancer cells in vitro, so this is believed to have the potential for the treatment of colorectal and other cancers. A deficiency in glucocerebrosidase activity may predispose individuals to more common disorders such as Parkinson's disease and Lewy body dementia. Excess glucosylceramide production and thence of more complex glycosphingolipids is a factor in polycystic kidney disease. It appears to be a general rule that the mere process of lysosomal substrate accumulation in all lysosomal storage disorders impairs lysosome integrity and results in more general disruptions to lipid metabolism and membrane structure and function. On the other hand, inhibition of glucosylceramidases may be of benefit in cystic fibrosis. Krabbe disease is discussed in the next section. Galactosylceramide is believed to function as an initial receptor for the human immunodeficiency virus (HIV) in mucosal epithelial cells and controls the early infection-independent phase of HIV transfer to T cells. Glucosylceramide levels regulate the uptake of viruses that rely upon the late endosomal compartment for fusion, including the influenza A and Ebola viruses. Gangliosides The name ganglioside was first applied by the German scientist Ernst Klenk in 1942 to a mixture of complex glycosphingolipids newly isolated from ganglion cells of brain. Subsequently, he demonstrated that as part of an oligosaccharide chain, they contained an acidic carbohydrate component, which he named "neuraminic acid" - later termed "sialic acid" from the Greek "sialon" for saliva, from which they were first isolated. However, it was not until 1963 that the first ganglioside species was fully characterized. Innumerable sphingolipids are now known that differ in the nature of both the glycan (glucose, galactose, N-acetylgalactosamine, and sialic acid residues) and ceramide structures. They are present throughout the animal kingdom, from echinoderms up to higher animals, but not in plants. Such highly polar, acidic and relatively hydrophilic molecules have distinctive physical properties, which are essential for the vital functions of gangliosides in the membranes of the central nervous system and other tissues. Sialic acids and Gangliosides Sialic acids: Gangliosides are oligoglycosylceramides derived as a first step from lactosylceramide, and they are defined by the presence of one to as many as five sialic acid residues, i.e. carbohydrate molecules with a nine-carbon backbone and a carboxylic acid group, a subclass of the superfamily of naturally occurring non‑2‑ulosonic acids. Of the many forms that have been characterized, only a few are relevant to gangliosides, and the most important of these is N-acetylneuraminic acid (‘NANA’ or ‘SA’ or 'Neu5Ac' or 'NeuAc'). Less often the sialic acid component is N-glycolylneuraminic acid (Neu5Gc), which differs by only one oxygen atom at the C-5 N-acetyl group, or it can be a Neu5Ac analogue in which the amide group is replaced by a hydroxyl group, i.e. 3-deoxy-D-glycero-D-galacto-nonulosonic acid (ketodeoxynonulosonic acid or ‘KDN’). The sialic acids are joined via α-glycosidic linkages to one or more of the monosaccharide units, e.g. via the hydroxyl group on position 2, or to another sialic acid residue. The polar head groups of the lipids carry a net-negative charge at pH 7.0 and they are acidic. Their structures are shown in Figure \(27\). Humans lack Neu5Gc: Neu5Ac is the biosynthetic precursor of Neu5Gc, a component of gangliosides from most animal species, including mice, horse, sheep, and goats, via the action of the enzyme CMP–N-acetylneuraminic acid hydroxylase (CMAH). However, NeuGc is not synthesized in humans (or birds and New World monkeys), although it is present in other primates such as the great apes, and indeed as it is a xeno-antigen, anti-NeuGc antibodies are produced normally in healthy humans (and especially after injection of NeuGc-containing glycoconjugates). The absence or irreversible inactivation of a number of relevant genes, but especially a critical exon in the CMAH gene, both for sialolipids and peptides in humans suggests that this may have been a major biochemical branch-point in human evolution that occurred ~2 to 3 million years ago after the divergence of humans and chimpanzees from a common ancestor. It may even be a factor in the superior performance of the human brain as the overexpression of Neu5Gc in the brains of transgenic mice was found to result in abnormal development. It could also mean that there might have been a fertility barrier between us and other hominids during evolution. While these are speculations, there is some evidence that the loss of Neu5Gc in humans had complex effects on immunity, providing greater capabilities to clear sublethal bacterial challenges. Some NeuGc may be obtained from the diet in meat and milk, for example, and this may be incorporated into human gangliosides to a limited extent, especially in fetal tissues and some cancers. In the latter, preferential expression of dietary Neu5Gc has been ascribed to their higher metabolic rate. 2. Structure and Occurrence of Gangliosides Most of the common range of gangliosides are derived from the ganglio- and neolacto-series of neutral oligoglycosphingolipids (Table 1), and they should be named systematically in the same way with the position of the sialic acid residue(s) indicated as for branched structures. However, they are more conveniently defined by a short-hand nomenclature system proposed by Svennerholm in which M, D, T and Q refer to mono-, di-, tri- and tetrasialogangliosides, respectively, and the numbers 1, 2, 3, etc refer to the order of migration of the gangliosides on thin-layer chromatography. For example, the order of migration of monosialogangliosides is GM3 > GM2 > GM1 (sometimes defined by subscripts, e.g. GM1 or GM1). To indicate variations within the basic structures, further terms are added, e.g. GM1a, GD1b, etc. Although alternatives have been proposed that are more systematic in structural terms, the Svennerholm nomenclature is that approved by IUPAC-IUB. Ganglio-series glycosphingolipids having 0, 1, 2 and 3 sialic acid residues linked to the inner galactose unit are termed asialo- (or 0-), a-, b- and c-series gangliosides, respectively, while gangliosides having sialic acid residues linked to the inner N-galactosamine residue are classified as α-series gangliosides. The structures for these groups are illustrated in the section on ganglioside biosynthesis below, for reasons of practical convenience. As of 2020, more than 200 gangliosides with variations in the carbohydrate chain had been characterized in vertebrates alone. One of the most studied monosialo-gangliosides and the first to be fully characterized is ganglioside GM1a (Neu5Acα2-3(Galβ1-3GalNAcβ1-4)Galβ1-4Glcβ1Cer), a major brain ganglioside of mammals and the preferred ligand of cholera toxin, illustrated in Figure \(28\). It can also be depicted using the abbreviated structure shown in Figure \(29\). An alternative nomenclature, which is less used, is recommended by IUPAC-IUB and is based upon the ganglio (Gg) root structure; it employs Roman numerals to designate each hexose unit and the location of the Neu5Ac along the carbohydrate chain with Arabic superscripts to designate the hydroxyl group to which this is linked. By this system, GM1a is defined as II3-α-Neu5Ac-Gg4Cer. Brain gangliosides: Gangliosides can amount to 6% of the weight of lipids from the brain (20 to 500 times more than in other tissues), where they constitute 10 to 12% of the total lipid content (20-25% of the outer layer) of neuronal membranes, for example. Aside from this, they are synthesized and are present at low levels (1 to 2% of the total lipids) in all animal tissues, where like the neutral oligoglycosphingolipids they are concentrated in the outer leaflet of the plasma membrane in the nanodomains known as 'rafts' or in related structures. Mammalian neurons actively synthesize gangliosides of the ganglio-series primarily, but oligodendrocytes in the brain produce instead myelin-forming glycosphingolipids, such as galactosylceramide and sulfatide together with a minor amount of ganglioside GM4. The brain contains as much as 20 to 500 times more gangliosides than most non-neural tissues, with three times as much in grey as in white matter. As the brain develops, there is an increase in the content of gangliosides and in their degree of sialylation. There are large differences between species and tissues. For example, during embryogenesis and the postnatal period in the human central nervous system, the total amount of gangliosides increases approximately threefold, while that of GM1 and GD1a increases 12 to 15-fold. During the same period, the hemato-series gangliosides GM3, GD3, and 9-OAc-GD3, which lack a hexosamine residue, are the predominant ganglioside species, but they are present in much lower amounts in adults and then in some areas of the brain only. In the mouse brain, the total amount of gangliosides is almost 8-fold greater in adults than in embryos, with a similar shift in composition from simple (GM3 and GD3) to more complex gangliosides. It is evident that the ganglioside changes during brain maturation are correlated with many neuro-developmental milestones, and there is no doubt that gangliosides play a crucial role in neuronal function and brain development, especially during infancy when there is high nutrient demand as the brain undergoes rapid restructuring. The main gangliosides (~95%) of adult mammalian brain are ganglio series GM1, GD1a, GD1b, and GQ1b, while lactosyl series gangliosides such as GM3 (sialyllactosylceramide) are found mainly in the extra-neural tissues. The remaining ~5% consists of minor components in the brain include gangliosides GM4, GM3, GD3, GM2, GD2, Fuc-GM1, Fuc-GD1b, GT1a and GP1c, the proportions of which vary depending on species. On the other hand, modern mass spectrometric methodology (electrospray ionization ion mobility MS) has revealed a much higher degree of sialylation than was previously recognized, including a complete series of mono- to octasialylated gangliosides in fetal frontal lobe. Subsequently, many previously unknown acetylated gangliosides were found in fetal hippocampus by this methodology. The content and composition of gangliosides in the brain also change with aging, with a substantial fall in the content of lipid-bound sialic acid but an increase in the proportion of the more complex forms in terms of carbohydrate structures in the elderly. Gangliosides in other tissues and species: Among the extraneural tissues, lactosyl series gangliosides such as GM3 (sialyllactosylceramide) and monosialogangliosides, in general, tend to predominate. Relatively high concentrations of ganglioside GD1a are present in erythrocytes, bone marrow, testis, spleen, and liver, while GM4 is more abundant in kidney, GM2 in bone marrow, GM1 in erythrocytes and GM3 in intestine. In germ cells of mice, there is a switch between gangliosides of the a- and 0-series upon differentiation when they are crossing the blood-testis barrier. Skin fibroblasts and many cells of visceral organs generate gangliosides of the globo series mainly. Similarly, glob-o and lacto series gangliosides are characteristic components of the stage-specific embryonic antigens (SSEA), which underlie the development and differentiation of human embryonic stem cells. A sialyl-lactotetraosylceramide is present in the latter and in the brains of children under the age of two, but not in tissues of adult humans. Gangliosides can cross the placental barrier into the fetus and those in milk, derived from the apical plasma membrane of secretory cells of the mammary gland, may be of nutritional importance for the newborn. GD3 is the main ganglioside in human breast milk at an early stage of lactation, whereas GM3 is more abundant in the later stages (and in bovine milk). Unfortunately, gangliosides are poorly characterized and quantified in foods in general. A 5-N-deacetylated form of ganglioside GM3 has been detected in human melanoma tumors. In addition, O-acetylation or lactonization of the sialic acid residue adds to the potential complexity. Gangliosides containing O-acetylated sialic acids, such as 9-OAc-GD3, are expressed during embryonic development and in the retina and cerebellum of adult rats, but not other brain regions. They occur also in certain tumors and may protect them from apoptosis. It is possible that such gangliosides are even more widespread, but they are missed after treatment with mild alkali during the isolation procedure, a common analytical practice. A further complexity is the occurrence of gangliosides with sulfate groups, and these have been isolated from human, mouse, and monkey kidney cells. KDN-containing gangliosides are minor components of egg, ovarian fluid, sperm and testis of fish and of some mammalian tissues Gangliosides from marine invertebrates (echinoderms), such as starfish and sea cucumbers, are very different in structure from those in vertebrates and do not have a shorthand nomenclature. They include forms with distinctive ceramide compositions, untypical carbohydrate residues, sialic acids within the oligosaccharide chain, or with glycosyl inositol-phosphoceramide structures. The mollusc, Aplysia kurodai, lacks gangliosides but produces complex oligoglycosylceramides with 2-aminoethylphosphonic acids and/or phosphoethanolamine groups attached that may serve as ganglioside surrogates. Ceramide structures: In general, the ceramide structures of gangliosides tend to be relatively simple. Sphingosine is usually the main sphingoid base, accompanied by the C20 analog in gangliosides of the central nervous system. Stearic acid (18:0) can be 80 to 90% of the fatty acid constituents in the brain, accompanied by small amounts of 16:0, 20:0 and 22:0, but with little or no polyunsaturated or 2-hydroxy acids, other than in some exceptional circumstances (e.g. some carcinomas). Palmitic acid is more abundant in gangliosides of the intestines and liver, while 2-hydroxylated fatty acids are relatively abundant in the last and in the kidney. There are also differences in the composition of the base and fatty acid components in different cells or regions of the brain. During development, the nature and concentrations of these constituents change markedly, and for example, the ratio of C20/C18-sphingosine in ganglioside GD1a of cerebellum increases 16-fold from 8-day-old to 2-year-old rats. In gangliosides outwith the nervous system, C20-sphingosine is barely detectable, and there is often a much wider range of fatty acid constituents (C14 to C24). The nature of the ceramide component is relevant to the biological function of gangliosides, and changing the fatty acid component to α-linolenic acid by synthetic means alters the biological activity of gangliosides dramatically in vitro. However, it is the carbohydrate moiety that has the primary importance for most of their functions, and detailed discussion of these structures would take us into realms of chemistry best left to carbohydrate experts (see the reading list below). In any given cell type, the number of different gangliosides may be relatively small, but their nature and compositions may be characteristic and in some way related to the function of the cell. It is noteworthy that some terminal glycan structures of gangliosides are also present in glycoproteins of membranes. 3. Biosynthesis There is evidence that the pool of glucosylceramide and thence of lactosylceramide that is utilized for ganglioside biosynthesis is different from that for the other neutral oligoglycosylceramides. This may explain some of the differences between the two groups in the fatty acid and sphingoid base components, which will also be dependent upon cell type. It is an open question how the ganglioside precursors enter the Golgi and trans-Golgi network where synthesis occurs at the luminal leaflet, but it appears that the regulation of intracellular sphingolipid traffic may be as important as the control of enzyme expression and activity in determining the final compositions of the various glycosphingolipid types. In humans, sialic acid biosynthesis occurs by a series of reactions in the cytosol, but the Neu5Ac produced is transferred to the nucleus and activated by the cytosine 5'-monophosphate N-acetylneuraminic acid synthetase (CMAS) to form CMP-Neu5Ac, which is transported to the Golgi apparatus by a family of sialyltransferases specific for particular glycosidic linkages (α2,3, α2,6, α2,8, and α2,9). Thereafter, the pathways for the biosynthesis of the common series of gangliosides of the ganglio-series, for example, involve sequential activities of distinct membrane-spanning sialyltransferases and glycosyltransferases as illustrated in Figure \(29\) for the four main 0-, a-, b- and c-series of gangliosides. The required enzymes are bound to the membranes of the Golgi apparatus in a sequence that corresponds to the order of addition of the various carbohydrate components. Thus, the sialyltransferase that catalyzes the synthesis of the relatively simple ganglioside GM3 is located in the cis-region of the Golgi, while those that catalyse the terminal steps of ganglioside synthesis are located in the distal or trans-Golgi region. The GM3 synthase in particular, which catalyzes the transfer of Neu5Ac from cytidine monophosphate (CMP)-Neu5Ac onto the terminal galactose residue of lactosylceramide, has a unique specificity. The simple ganglioside GM3 is synthesized by the addition of sialic acid to lactosylceramide by CMP:LacCer α2-3 sialyltransferase (or GM3 synthase), before GD3 and GT3 are produced in turn by the action of appropriate synthases. Subsequently, GM3, GD3 and GT3 serve as precursors of more complex gangliosides by the action of further glycosyl- and sialyl-transferases. An alternative theory with some supporting evidence proposes that a multiglycosyl-transferase complex is responsible for the synthesis of each individual ganglioside rather than a series of individual enzymes. Further sialylation of each of the a, b, and c series and in different positions in the carbohydrate chain can occur to give an increasingly complex and heterogeneous range of products, such as the α-series gangliosides with sialic acid residue(s) linked to the inner N-acetylgalactosamine residue (not illustrated). GM4 or NeuAcα2,3Gal-Ceramide, a minor component of the brain and present in a few other tissues at low levels, is an exception in that galactosylceramide is its precursor. Finally, the newly synthesized gangliosides are transferred to the external leaflet of the plasma membrane via the lumenal surface of transport vesicles. Gangliosides are also important constituents of nuclear membranes. The changes that occur in ganglioside compositions of brain and other tissues in the embryonic and post-natal stages are governed mainly by changes in the expression level and activity of the glycosyl- and sialyl-transferases, although the former can also be regulated by glycosylation and phosphorylation. The presence of distinctive sialidases that differ from the catabolic lysosomal enzymes (see below) in raft-like regions of the plasma membrane bring about further changes in the composition of the cell surface gangliosides that can be specific to particular cell types, causing a shift from poly-sialylated species involving a decrease of GM3 and formation of GM2 then GM1 by hydrolysis of terminal sialosyl residues linked either α2‑8 on another sialic acid or α2‑3 on galactose. As GM1 is resistant to most sialidases, it tends to increase in concentration relative to oligosialo species as developmental and other GM1-requiring processes come into play. This may have consequences for important cellular events, such as neuronal differentiation and apoptosis. Conversely, sialylation may occur in some neuronal membranes, increasing the proportions of poly-sialylated species. In particular, a CMP-NeuAc:GM3 sialyltransferase is able to sialylate GM3. Gangliosides GM1 and GD1a have been identified in both membranes of the nuclear envelope together with two neuraminidases. Ganglioside lactones, where the sialic acids are linked together with ester linkages, have been detected as minor components in brain tissues, where lactonization occurs at the plasma membrane. As the process of lactonization profoundly influences the shape and biological properties of the original ganglioside, it is possible that lactonization-delactonization in a membrane might be a trigger for specific cellular reactions. Similarly, GD3 ganglioside can undergo O-acetylation at C9 of the outer sialic acid with important metabolic implications. Gangliosides added to many types of cell preparations in vitro are rapidly taken up by the cells, while gangliosides injected into animals in vivo are rapidly internalized by tissues. They can cross the blood-brain barrier, and via the placenta, they can enter the fetus. Similarly, dietary gangliosides are absorbed intact by intestinal cells but are broken down to their lipid and carbohydrate constituents for re-use. The sialic acids released by an intestinal sialidase are transported in plasma to the brain and other tissues where they influence ganglioside expression. Indeed, there is some experimental evidence that dietary gangliosides may improve cognitive functions in animals and humans. Catabolism Degradation of gangliosides takes place at the surface of intralysosomal luminal vesicles, generated by an inward budding of the endosomal membrane, and these are reached by a process of endocytosis. In brief in relation to gangliosides, soluble sialidases (neuraminidases) and exoglycohydrolases remove individual sialic acid and sugar residues sequentially from the non-reducing terminal unit, as illustrated for ganglioside GM1, with the eventual formation of ceramide, which is then split into long-chain base and fatty acids by ceramidases. This degradation occurs through the endocytosis-endosome-lysosome pathway with a requirement for an acidic pH inside the organelle. In addition to the sialidases and exoglycohydrolases, the various reactions have an absolute requirement for effector molecules, termed 'sphingolipid activator proteins', including saposins (Sap), and the specific GM2-activator protein (GM2-AP). Ganglioside GM3 is a component of the lysosomal perimeter membrane, but is protected from degradation by a glycocalix of the membrane facing the lysosol. Anionic lipids and especially bis(monoacylglycero)phosphate in the membranes stimulate ganglioside degradation while cholesterol is inhibitory. The catabolic pathway is shown in Figure \(30\). This process constitutes a salvage mechanism that is important to the overall cellular economy since a high proportion of the various hydrolysis products are recycled for glycolipid biosynthesis. By generating ceramide and sphingosine, it may also be relevant to the regulatory and signaling functions of these lipids. In addition, some partial hydrolysis of gangliosides occurs in the plasma membrane as part of a biosynthetic remodeling process discussed above. Defects in catabolism lead to the gangliosidoses discussed later. Ganglioside Function Cell surface effects: In their natural biological environment, gangliosides have a negative charge because of the presence of sialic acids, which also add to the hydrophilicity of the polysaccharide constituent. This is balanced somewhat by the hydrophobic character of the ceramide moiety, so that over all the molecules are amphiphilic in nature, but very different from the glycerophospholipids, which are essential for the formation of membrane bilayers. Indeed, a ganglioside such as GM1 is virtually soluble in water, where it can form large aggregates though hydrophilic effects. The nature of the ceramide unit with its capacity to form hydrogen bonds with glycerophospholipids is important in ensuring that gangliosides are inserted in a stable manner into the outer layer of the plasma membrane. Thus, gangliosides are anchored in membranes by their ceramide units with the double-tailed sialoglycan components extending out from the cell surface, where they can participate in intermolecular interactions by a network of hydrogen bonds and hydrophobic interactions. For example, the glucose-ceramide bond of GM1 is oriented in the outer leaflet of the plasma membrane such that the glycan extends perpendicularly to the plane of the lipid bilayer. All gangliosides, but especially the simplest GM3 or Neu5Acα2-3Galβ1-4Glcβ1Cer, have a structural role, and they a natural propensity to laterally segregate and to associate with each other and with other sphingolipids, phospholipids and cholesterol into raft nano-domains or in related structures, such as the caveolae, where the very large surface area occupied by the oligosaccharide chain imparts a strong positive curvature to the membrane. In this environment, gangliosides can interact with each other through side-by-side hydrogen bonds mediated by water molecules that act as bridges between the chains. Further, molecules of GM3 and other gangliosides self-aggregate into clusters on the surface of lymphocytes of human peripheral blood, and there is evidence that the density of these clusters in membranes governs their reactivity as antigens. In addition, it is believed that gangliosides and other oligoglycosylceramides can cluster together through hydrogen donor-acceptor (cis) interactions because of the presence of hydroxyl and acetamide groups to form glycosynaptic domains, which are related to but functionally distinct from raft signaling platforms (with lower cholesterol concentrations). Many of the biological functions of gangliosides are mediated through their location in these nanodomains, where they may have specialized functions in cell adhesion, growth, and motility through interactions with specific proteins and signal transduction pathways. However, not all gangliosides are present in such raft-like structures. Receptor/signaling functions: Gangliosides can bind to membrane proteins directly by carbohydrate-carbohydrate or carbohydrate-amino acid interactions, usually involving specific ganglioside head groups, resulting in changes to the location of proteins within membrane microdomains for recruitment of signaling partners, or to dimerization or other effects upon receptors. In rafts and caveolae especially, gangliosides can modulate cell signaling processes by their interactions with specific receptors, adhesion molecules, and ion channels. Cell–cell (trans) interactions occur by sialoglycans on one cell binding to complementary binding proteins (lectins) on adjacent cells, bringing about adhesion of cells and enabling regulation of intracellular signaling pathways, e.g. myelin-associated glycoprotein on myelin sheaths binds to gangliosides present on axonal membranes. In addition, gangliosides act as receptors of interferon, epidermal growth factor, nerve growth facto,r and insulin, and they may regulate cell signaling and control growth and differentiation of cells in this way. While intact gangliosides inhibit growth by rendering cells less sensitive to stimulation by epidermal growth factor, removal of the N-acetyl group of sialic acid enhances this reaction and stimulates growth. Gangliosides function as antigens or receptors by recognizing specific molecules (lectins), including bacterial toxins, at the cell surface and by modulating the charge density at the membrane surface (see the section on Gangliosides and Disease below). They also regulate the activities of proteins within the plasma membrane and especially receptor-type tyrosine kinases. For example, the phosphorylation state and activity of insulin receptors in caveolae and thence the insulin resistance of cells is controlled by the concentration of GM3, the main ganglioside in plasma and other extraneural tissues. GM3 interacts also with the epidermal growth factor receptor leading to cell growth inhibition. GM1 strongly influences specific neuronal functions by interacting with specific receptors such as the tropomyosin receptor kinase (Trk) A (TrkA) receptor by altering its conformation to enable interaction with the nerve growth factor (NGF) ligand. GM3 (SA-Gal-Glc-Cer) is a serum ganglioside that is highly enriched in a type of membrane microdomain termed a 'glycosynapse', and it forms complexes with co-localized cell signaling molecules. It has a function in the innate immune function of macrophages and it has been demonstrated that molecular species of GM3 with differing acyl-chain structures and modifications can operate as pro- and anti-inflammatory modulators of Toll-like receptor 4 (TLR4); very-long-chain and α-hydroxy GM3 species increase TLR4 activation, while long-chain and unsaturated GM3 species have the opposite effect. In addition, gangliosides have been shown to be cell-type specific antigens that have key functions in immune defense. For example, a major immunological function of gangliosides and sialic acids is to protect cells from attack by our own immune system and from autoimmunity. They recognize and protect host organs and tissues from complement attack by binding to the complement regulatory protein factor H, which has the potential to exert strong cytotoxic and inflammation-inducing activity. In particular, sialic acids protect against complement killing of autologous cells by binding to this protein via the α2–3 linked sialic acid glycans of the GD3 ganglioside. On the other hand, the breakdown of this system can lead to autoimmune diseases. Brain function: One of the first examples of a ganglioside influencing a signaling event to be studied in some detail concerns the simple ganglioside GD3, which has a central role in early neurogenesis. GD3 binds to the epidermal growth factor receptor (EGFR) via a protein-carbohydrate interaction involving its terminal N-acetylneuraminic acid and a lysine residue in the transmembrane domain of the receptor and also by a carbohydrate-carbohydrate interaction thereby maintaining the latter in its inactive monomeric state. EGFR then binds to the epidermal growth factor and stimulates the transition of the receptor from an inactive monomeric to an active homodimeric form, and this in turn triggers receptor auto-phosphorylation and activation of a signaling cascade that promotes cell proliferation. This has proven to be essential for the regulation of the stem cell self-renewal capacity in the brain. In contrast, the neutral oligoglycosphingolipid Gb4 exerts the opposite effect on EGFR by interacting directly with it to potentiate its auto-phosphorylation with activation of the downstream cascade. The techniques of molecular biology such as targeted gene deletion, which enable specific enzymes to be eliminated from experimental animals, are now leading to a better understanding of the function of each ganglioside. It is evident that they are essential to central myelination, to maintain the integrity of axons and myelin, and for the transmission of nervous impulses. These effects may be mediated by interactions of the negatively charged sialic acid residues of gangliosides with calcium ions, which are critical for neuronal responses. For example, a variant of GD3, 9-O-acetyl GD3, appears to be involved in glial-guided neuronal migration during brain development in the rat, while GM1 may have a similar function in humans; it determines which growth cone of unpolarized neurons becomes the axon. By stabilizing neuronal circuits, gangliosides have a function in memory, and conversely, disturbances in ganglioside synthesis can lead to neurodegenerative disorders (see below). Ganglioside GM3 in raft domains has been shown to have an indispensable role for the development, function, and viability of cochlear hair cells and thence it is essential for hearing. On the other hand, mice that express GM3 primarily and are devoid of the typical complex gangliosides of the brain suffer weight loss, progressive motor and sensory dysfunction, and deterioration in spatial learning and memory with aging. GD3 is important for retinal structure and visual function in mice. Changes in ganglioside composition can be induced by nerve stimulation, environmental factors, or drug treatments. The various interconvertible ganglioside types in the plasma membrane of neurons are particularly important for its development in that they regulate such processes as axonal determination and growth, signaling, and repair. In addition, gangliosides are believed to be functional ligands for the maintenance of myelin stability and the control of nerve regeneration by binding to a specific myelin-associated glycoprotein. The occurrence of gangliosides in cell nuclei suggests a possible involvement of gangliosides in the expression of genes relevant to neuronal function. For example, the monosialoganglioside GM1 has been shown to promote the differentiation of various neuronal cell lines in culture. It has protective effects on the neural system by encouraging neural stem cell survival and proliferation, while facilitating the stability and regeneration of axons, and by inhibiting neurodegeneration through autophagy, for example after ischemic stroke. Within membrane rafts, this ganglioside has key roles in several signaling systems through association with specific proteins that have glycolipid-binding domains, including those that modulate mechanisms such as ion transport, neuronal differentiation, G protein-coupled receptors (GPCRs), immune system reactivities and neuroprotection. It is important for Ca2+ and Na+ homeostasis in the nucleus and plasma membrane and in regulating the effects of platelet-derived growth factor. However, there have been unpleasant complications when GM1 has been administered for therapeutic purposes. GD1a is sometimes considered to be a reserve pool for GM1. After nerve injury, toll-like receptor 2 (TLR2) signaling is important for the induction of neuropathic pain; ganglioside GT1b functions as a TLR2 agonist to produce mechanical and thermal hypersensitivity. Other functions: The ganglioside GD3 is essential for the process of apoptosis by blocking the activation of specific transcription factors and thence disabling the induction of antiapoptotic genes. 9-O-Acetylation of the GD3 molecule prevents ganglioside oxidation and blocks its pro-apoptotic effects. Similarly, GD3 is a regulator of autophagy, i.e. the degradation and/or recycling of cellular components. Gangliosides are also important in reproduction, and in mice, GD1a has been shown to be important to oocyte maturation, monospermic fertilization, and embryonic development, while GM1 is important in sperm-oocyte interactions and sperm maturation processes. Deletion of the GM2/GD2 synthase leads to infertility in male mice and the production of a novel fucosylated ganglioside containing very-long-chain polyunsaturated fatty acids. Related studies with gene knockout mice have revealed that b-series gangliosides are important in leptin secretion from adipocytes, while a-series gangliosides interact with the leptin receptor in the hypothalamus to influence the balance of energy. Gangliosides and Disease Bacterial toxins and viruses: In relation to adaptive immunity, a-series and o-series gangliosides in the plasma membrane are involved in the function and stimulation of receptors on certain subsets of T cells by acting as pattern-recognition receptors for invading pathogens. In particular, certain gangliosides bind specifically to viruses and to various bacterial toxins, such as those from botulinum, tetanus and cholera, and to blood merozoites of the deadliest malaria parasite Plasmodium falciparum, and they mediate interactions between microbes and host cells during infections, with NeuAc as the main recognition module. The best known example is cholera toxin, which is an enterotoxin produced by Vibrio cholerae where the specific cell surface receptor is ganglioside GM1; the five B-chains of cholera toxin each bind one molecule of GM1. Interestingly, the subsequent metabolism of the ganglioside-toxin complex is dependent on the nature of the fatty acid components of the ganglioside. It is believed that toxins utilize the gangliosides to hijack an existing retrograde transport pathway from the plasma membrane to the endoplasmic reticulum. For example, the passage of the cholera toxin through the epithelial barrier of the intestine is mediated by GM1, possibly by endocytosis of the toxin-GM1 complex via caveolae into the apical endosome and thence into the Golgi/endoplasmic reticulum, where the complex dissociates. The consequence is persistent activation of adenylate cyclase by the toxin and continuous production of cAMP that leads to the severe fluid loss typical of cholera infections. As a further example, the botulinus toxin binds to a complex of a polysialoganglioside with the protein synaptotagmin, which together act as a high-affinity receptor complex to enable the neurotoxic effects. Similarly, ganglioside GM2 binds to a toxin secreted by Clostridium perfringens. Influenza viruses have two glycoproteins in their envelope membranes, hemagglutinins, which bind to cellular receptors such as gangliosides, and after entry into respiratory epithelial cells, the sialidase (neuraminidase) of the virus cleaves the sialic acid from the receptors to prevent entry of further viruses to the cell. Variations in the structure of these proteins force the development of new vaccines The carbohydrate moiety of gangliosides is essential for the initial binding of viruses, but the lipid moiety is believed to be important for controlling their intracellular transport. Some gangliosides and GD1a especially have anti-inflammatory properties in that they inhibit the effects of bacterial lipopolysaccharides by preventing the activation of tumor necrosis factor (TNF) and other cytokines. In contrast, GM2 may increase cytokine production in similar circumstances, while the heat-labile toxins of Escherichia coli bind to several gangliosides in macrophages, thus activating an inflammatory response. Gangliosidoses and other neurodegenerative diseases: It appears to be a general rule that the mere process of lysosomal substrate accumulation in all lysosomal storage disorders impairs lysosome integrity and results in more general disruptions to lipid metabolism and membrane structure and function, inevitably triggering pathologic mechanisms. Endogenous generation of antibodies to gangliosides is often a factor, and it has been argued that gangliosides and their sialic acids components are at the border of immune tolerance. As with the neutral oligoglycosylceramides and ceramide monohexosides, a number of unpleasant lipidoses have been identified that involve the storage of excessive amounts of gangliosides in tissues because of failures in the catabolic mechanism. The most important of these are the GM2 gangliosidoses, i.e. Tay-Sachs disease (and the similar Sandhoff disease), a fatal genetic disorder found mainly in Jewish populations in which harmful quantities of ganglioside GM2 accumulate in the nerve cells in the brain and other tissues. Lyso-GM2 (non-acylated) in plasma may serve as a marker. A modified GM2 derivative that contains taurine in amide linkage to the sialic acid carboxyl group has been identified in the brain of such patients. As infants with the most common form of the disease develop, the nerve cells become distended and a relentless deterioration of mental and physical abilities occurs. The condition is caused by insufficient activity of specific enzymes, i.e. β‑N‑acetylhexosaminidase, which catalyzes the degradation of gangliosides by removing the terminal N-acetylgalactosamine residue from GM2, or the GM2 activator protein. In addition, a generalized GM1 gangliosidosis (an autosomal recessive and neurodegenerative disease) has been characterized in which ganglioside GM1 accumulates in the nervous system leading to mental retardation and enlargement of the liver. The condition is a consequence of a deficiency of the lysosomal β-galactosidase enzyme, which hydrolyses the terminal β-galactosyl residues from GM1 ganglioside to produce GM2. It appears that storage of substantial amounts of unwanted lipids in the lysosomal system leads to a state of cellular starvation, so that essential elements such as iron are depleted in brain tissue. The presence of lyso-GM1 in plasma is now seen as a useful aid to diagnosis. Small amounts of some gangliosides accumulate as secondary storage compounds in Niemann–Pick disease. The Guillain–Barré syndrome is an acute inflammatory disorder, usually triggered by a severe infection, which affects the peripheral nervous system. Antibodies to gangliosides are produced by the immune system, leading to damage of the axons, which can result in paralysis of the patient. Huntington’s disease is believed to involve disruption of the metabolic pathways between glycosylceramides and gangliosides, and there is a human autosomal recessive infantile-onset epilepsy syndrome caused by a mutation to a sialyl transferase. Impaired ganglioside metabolism is also relevant to Alzheimer’s disease, because complexation with ganglioside GM1 may cause aggregation of the amyloid β-protein deposits that characteristically accumulate in the brain in this condition (this explanation does not appear to be universally accepted). In general, in ganglioside deficiencies, natural or induced, it appears that progressive inflammatory reactions take place, leading to neurodegeneration in part because of the deterioration of the architecture of lipid rafts. On the other hand at normal tissue concentrations, gangliosides such as GM1 are believed to have an anti-inflammatory and neuroprotective role in certain types of neuronal injury, Parkinsonism, and some related diseases. For example in relation to Parkinson's disease, GM1 binds to α-synuclein and inhibits or eliminates fibril formation. It may have a protective role by preventing sphingomyelin-induced aggregation, although as the overall level of GM1 decreases during aging, its beneficial effect decreases. For these reasons, the therapeutic properties of ganglioside GM1, the most accessible species, and derived molecules are under clinical investigation. However, there is no approved therapy for any gangliosidosis, although a number of different therapeutic strategies are being studied, including hematopoietic stem cell transplantation and gene therapy. For the moment, the blood-brain barrier remains a challenge. Cancer: Gangliosides have important functions in cancer, especially in the regulation of signal transduction induced by growth-factor receptors in a specific microdomain termed a 'glycosynapse' in the cancer cell membranes, and in interactions with glycan recognition molecules involved in cell adhesion and immune regulation. In particular, depending on tissue, certain distinctive gangliosides are expressed at much higher levels in tumors than in normal healthy tissues, mainly by aberrant expression of glycosyltransferases and glycohydrolases. This enables tumor cells to escape immune surveillance and retain their malignancy. GM3 is not expressed in melanocytes normally, but is detected in 60% of primary melanomas and in 75% of metastatic melanomas, for example. Gangliosides can be shed from the surface of tumor cells into the local environment where they can influence interactions between cancer cells, including the transition of tumors from a dormant to a malignant state (angiogenesis); when present in the circulation they can be useful diagnostic aids. For example, the ganglioside GM3 is elevated in the serum of patients with breast cancer and may be a biomarker for the disease, while disialylated gangliosides GD2 and GD3 (Figure \(31\)) are considered to be markers of neuroectoderm origin in tumors (neuroblastoma). Specific gangliosides can have either positive or negative effects upon the regulation of the malignant properties of cancer cells. As a generality, disialyl glycosphingolipids or tandem-repeated sialic acid-structures confer malignant properties in various cancer systems; they are not merely markers. For example, the disialo-gangliosides GD2 and GD3 are present in trace amounts only in normal tissues, but are found at much higher concentrations in cancer cells, especially melanomas and neuroblastomas, with GD2 especially elevated in triple-negative breast cancer. These b-series gangliosides play a substantial part in the malignant properties of gliomas by mediating cell proliferation, migration, invasion, adhesion, and angiogenesis, and in preventing immunosuppression. They are considered to be tumor-associated antigens, and the GD2 and GD3 synthases are seen as important drug targets. In contrast, monosialyl gangliosides, such as GM1, GM2 and GM3, may suppress the malignant properties of various cancer cells. The mechanism is believed to involve complex formation at the cell surface with membrane proteins, such as growth factor receptors and adhesion receptors like those of the integrin family, leading to the modification of cell signals mediated by these receptors. Metastatic melanoma cells have high levels of GD3 in comparison to poorly metastatic cells or the normal counterpart, suggesting that GD3 may promote metastasis possibly by suppressing the anti-tumor immune response. Ganglioside GM3(Neu5Gc), i.e. containing an abnormal sialic acid, is sometimes considered to be a tumor-specific antigen and a target for cancer immunotherapy. Aberrant sialylation is found in many malignant cancers, where the levels of neuraminidases are key factors for metastasis and survival of cancer cells, and there can be a significant accumulation of unusual gangliosides containing N-glycolyl sialic acid in some cancers. N-Glycolyl-GM3, normally absent from human tissues, is present in all stage II breast cancers, and it is accompanied by a number of other less common complex gangliosides. Similarly, the 5-N-deacetylated form of GM3 is expressed in metastatic melanomas, but not in healthy tissue or even in primary melanomas; it is considered to be a specific marker for the metastatic condition and a target for potential therapy. Increased synthesis of 9-O-acetyl-GD3, dependent on a sialyl-O-acetyltransferase - CAS1 Domain-Containing Protein 1, occurs in acute lymphoblastic leukemia and in malignant melanomas, and this appears to limit apoptosis, while O-acetylated GD2 (OAcGD2) is expressed in breast cancer and other tumors. A unique fucosyl-GM1 in which the terminal galactose is α-1,2-fucosylated at the non-reducing end is found circulating in the serum of patients with a number of cancers and especially with small-cell lung cancer but rarely in normal conditions, and it is also considered to be a potential indicator of cancer and a candidate for immunotherapy. Clinical trials with an antibody to GD2 have been carried out successfully against the rare childhood cancer neuroblastoma, and the USDA has approved the use of this in combination with other drugs to treat this often lethal cancer. However, this antibody can have painful side effects due to an interaction with GD2 on neurons, and modified antibodies, which may be safer, are now being tested in multiple clinical trials. A phase I clinical trial with an antibody to GD3 has shown promising results in patients with malignant melanoma. Similarly, antibodies to OAcGD2 and fucosyl-GM1 have shown anti-tumour effects in vitro, and studies with human patients are underway. Other diseases: Aberrant production of the ganglioside GM3 has been linked to pathophysiological changes associated with obesity, metabolic disorders, and type 2 diabetes mellitus through its effects on insulin receptors. It has a role in autoimmune disorders such as multiple sclerosis. In epilepsy, it is believed that a deficiency in the enzyme ceramide synthase 1, which produces 18:0 ceramides, leads to reduced ganglioside formation. By their presence in certain subsets of T cells, gangliosides influence allergic responses and auto-immune diseases. As gangliosides are present on the surface of vascular, vascular-associated, and inflammatory cells, they may have a role in atherosclerosis and in aging.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/21%3A_Lipid_Biosynthesis/21.04%3A_Biosynthesis_of_Membrane_Sphingolipids.txt

Search Fundamentals of Biochemistry By William (Bill) W. Christie and Henry Jakubowski. This section is an abbreviated and modified version of material from the Lipid Web, an introduction to the chemistry and biochemistry of individual lipid classes, written by William Christie. Sterols: Cholesterol and Cholesterol Esters In animal tissues, cholesterol (cholest-5-en-3β-ol) is by far the most abundant member of a family of polycyclic compounds known as sterols. It can also be described as a polyisoprenoid or a triterpene from its biosynthetic origin. Cholesterol was first recognized as a component of gallstones as long ago as 1769, while the great French lipid chemist Chevreul isolated it from animal fats in 1815. However, it was well into the 20th century before the structure was fully defined by the German Chemist Heinrich Wieland, who received the Nobel Prize in Chemistry for his work in 1927, the first of thirteen so honored for research on cholesterol and its metabolism. Cholesterol plays a vital role in animal life, and it is essential for the normal functioning of cells both as a structural component of cell membranes and as a precursor of steroid hormones and other key metabolites including vitamin D and bile acids. It is also important for cell signaling, transport processes, nerve conduction and the regulation of gene transcription. Every cell in vertebrates is able both to synthesize cholesterol and to metabolize it, and there is evidence that synthesis de novo is essential whatever the dietary intake; this is vital in the brain. However, excess cholesterol can contribute to the pathology of various diseases, notably cardiovascular disease, so cholesterol levels must be balanced to ensure an adequate but not excessive supply. Cholesterol – Structure, Occurrence, and Function in Membranes The struture of cholesterol is shown below in Figure \(1\). Cholesterol consists of a tetracyclic cyclopenta[a]phenanthrene structure with an iso-octyl side-chain at carbon 17. The four rings (A, B, C, D) have trans ring junctions, and the side chain and two methyl groups (C-18 and C-19) are at an angle to the rings above the plane with β stereochemistry (as for the hydroxyl group on C-3 also); there is a double bond between carbons 5 and 6. Thus, the molecule has a rigid planar four-ring nucleus with a flexible tail. Of the two recognized numbering systems in use, one originally described by Fieser and Fieser in 1959 and a second by IUPAC-IUB in 1989, the first appears to be preferred by most current authors. Most of the atoms in cholesterol can be placed on the diamond lattice, which is the structure showing the position of each carbon atom in the network solid diamond. In that structure, each carbon is connected to four other carbons using sp3-hybridized atomic orbitals and tetrahedral geometry. A representation of the diamond lattice is shown below. It consists of a series of interconnected boat conformations of cyclohexane propagating in the xyz direction. The structures of two different fused cyclohexanes, trans- and cis-decalins as well as the structure of three fused cyclohexanes, adamantane, are shown in Figure \(2\). Superimposing sp3-connected atoms onto a diamond lattice allows improved visualization of the real and allowed structures of more complicated molecules. Figure \(3\) shows a superposition of 5-α-cholestane, a reduced and non-hydroxylated form of cholesterol, onto the diamond lattice. The pink cyclopentane D ring of 5-α-cholestane, which is distorted from the 1090  bond angle for sp3-hybridized carbon atoms, does not fit exactly onto the lattice. Cholesterol, with its double bond, would also deviate from the ideal position on the diamond lattice given the two sp2-hybridized carbons in the double bond. Cholesterol is a ubiquitous component of all animal tissues (and of some fungi), produced by every nucleated animal cell, where much of it is located in the membranes, although it is not evenly distributed. The highest proportion of unesterified cholesterol is in the plasma membrane (roughly 30-50% of the lipid in the membrane or 60-80% of the cholesterol in the cell), while mitochondria and the endoplasmic reticulum have much less (~5% in the latter), and the Golgi contains an intermediate amount. Cholesterol is also enriched in early and recycling endosomes, but not in late endosomes. It may surprise some to learn that the brain contains more cholesterol than any other organ, where it comprises roughly a quarter of the total free cholesterol in the human body, 70-80% of which is in the myelin sheath. Of all the organic constituents of blood, only glucose is present in a higher molar concentration than cholesterol. In animal tissues, it occurs in the free form, esterified to long-chain fatty acids (cholesterol esters), and in other covalent and non-covalent linkages, including an association with the plasma lipoproteins. In plants, it tends to be a minor component only of a complex mixture of structurally related 'phytosterols', although there are exceptions but it is nevertheless importance as a precursor of some plant hormones. Animals in general synthesize a high proportion of their cholesterol requirement, but they can also ingest and absorb appreciable amounts from foods. On the other hand, many invertebrates, including insects, crustaceans and some molluscs cannot synthesize cholesterol and must receive it from the diet; for example, spiny lobsters must obtain exogenous cholesterol to produce essential sex hormones. Similarly, it must be supplied from exogenous sources to the primitive nematode Caenorhabditis elegans, where it does not appear to have a major role in membrane structure, other than perhaps in the function of ion channels, although it is essential the production of steroidal hormones required for larval development; its uptake is regulated by the novel lipid phosphoethanolamine glucosylceramide. Some species are able to convert dietary plant sterols such as β-sitosterol to cholesterol. Prokaryotes lack cholesterol entirely with the exception of some pathogens that acquire it from eukaryotic hosts to ensure their intracellular survival (e.g., Borrelia sp.); bacterial hopanoids are often considered to be sterol surrogates. Cholesterol has vital structural roles in membranes and in lipid metabolism in general with an extraordinary diversity of biological roles, including cell signaling, morphogenesis, lipid digestion and absorption in the intestines, reproduction, stress responses, sodium and water balance, and calcium and phosphorus metabolism, and we can only touch on a few of these functions in this web page. It is a biosynthetic precursor of bile acids, vitamin D, and steroid hormones (glucocorticoids, estrogens, progesterones, androgens, and aldosterone), and it is found in covalent linkage to specific membrane proteins or proteolipids ('hedgehog' proteins), which have vital functions in embryonic development. In addition, it contributes substantially to the development and working of the central nervous system. On the other hand, excess cholesterol in cells can be toxic, and a complex web of enzymes is essential to maintain the optimum concentrations. Because plasma cholesterol levels can be a major contributory factor to atherogenesis, media coverage has created what has been termed a ‘cholesterophobia’ in the population at large. One of the main functions of cholesterol is to modulate the fluidity of membranes by interacting with their complex lipid components, specifically the phospholipids such as phosphatidylcholine and sphingomyelin. As an amphiphilic molecule, cholesterol is able to intercalate between phospholipids in lipid bilayers to span about half a bilayer. In its three-dimensional structure, it is in essence a planar molecule that can interact on both sides. The tetracyclic ring structure is compact and very rigid. In addition, the location of the hydroxyl group facilitates the orientation of the molecule in a membrane bilayer, while the positions of the methyl groups appear to maximize interactions with other lipid constituents. The structure of cholesterol as it would appear on the diamond lattice is shown in Figure \(4\). As the α-face of the cholesterol nucleus (facing down) is 'smooth', it can make good contact with the saturated fatty-acyl chains of phospholipids down to about their tenth methylene group; the β-face (facing up) is made 'rough' by the projection of methyl groups from carbons 10 and 13. The interaction is mainly via van der Waals and hydrophobic forces with a contribution from hydrogen bonding of the cholesterol hydroxyl group to the polar head group and interfacial regions of the phospholipids, especially sphingomyelin. Intercalated cholesterol may also disrupt electrostatic interactions between the ionic phosphocholine head groups of nearby membrane phospholipids, leading to increased mobility of the head groups. Indeed, there is evidence that cholesterol forms stoichiometric complexes with the saturated fatty acyl groups of sphingomyelin and to a lesser extent of phosphatidylcholine. Experiments with mutant cell lines and specific inhibitors of cholesterol biosynthesis suggest that an equatorial hydroxyl group at C-3 of sterols is essential for the growth of mammalian cells. The Δ5 double bond ensures that the molecule adopts a planar conformation, and this feature also appears to be essential for cell growth, as is the flexible iso-octyl side-chain. The C-18 methyl group is crucial for the proper orientation of the sterol. While plant sterols appear to be able to substitute for cholesterol in supporting many of the bulk properties of membranes in mammalian cells in vitro, cholesterol is essential for other purposes. In the absence of cholesterol, a membrane composed of unsaturated lipids is in a fluid state that is characterized by a substantial degree of lipid chain disorder, i.e., it constitutes a liquid-disordered phase. The function of cholesterol is to increase the degree of order (cohesion and packing) in membranes, leading to the formation of a liquid-ordered phase. In contrast, it renders bilayers composed of more saturated lipids, which would otherwise be in a solid gel state, more fluid. Thus, cholesterol is able to promote and stabilize a liquid-ordered phase over a substantial range of temperatures and sterol concentrations. Further, high cholesterol concentrations in membranes reduce their passive permeability to solutes. These effects enable membranes to bend or withstand mechanical stresses, and they permit the fine-tuning of membrane lipid composition and organization, and regulate critical cell functions. Simplistically, the higher cholesterol concentrations in the plasma membrane support its barrier function by increasing membrane thickness and reducing its permeability to small molecules. In contrast, the endoplasmic reticulum has increased membrane flexibility because of its lower cholesterol concentrations and thus enables the insertion and folding of proteins in its lipid bilayer. While mitochondrial membranes have a low cholesterol content in total, this may be concentrated in nanodomains at regions of high curvature in the inner mitochondrial membrane with links to nucleoprotein complexes (nucleoids). In comparison to other lipids, it has been reported that cholesterol can flip rapidly between the leaflets in a bilayer, although this does not appear to be accepted universally, leading to doubts as to the trans-bilayer distribution of cholesterol in some biological membranes. However, much recent evidence suggests that the concentration of cholesterol in the inner leaflet of the plasma membrane is much lower than that in the outer leaflet in a range of mammalian cells. This distribution is important in that cholesterol promotes negative curvature of membranes and may be a significant factor in bringing about membrane fusion in the process of exocytosis. It may also be relevant for the regulation of various cellular signaling processes at the plasma membrane. Cholesterol also has a key role in the lateral organization of membranes and their free volume distribution, factors permitting more intimate protein-cholesterol interactions that may regulate the activities of membrane proteins. Many membrane proteins bind strongly to cholesterol, including some that are involved in cellular cholesterol homeostasis or trafficking, and contain a conserved region termed the ‘sterol-sensing domain’. Some proteins bind to cholesterol deep within the hydrophobic core of the membrane via binding sites on the membrane-spanning surfaces or in cavities or pores in the proteins, driven by hydrogen bond formation. Cholesterol has an intimate interaction with G-protein-coupled receptors (GPCRs) to affect ligand binding and activation, either by direct high-affinity binding to the receptor, by changing their oligomerization state, or by inducing changes in the properties of the membrane. For example, it is essential for the stability and function of the β2-adrenergic, oxytocin and serotonin receptors by increasing the agonist affinities, while the inactive state of rhodopsin is stabilized both through indirect effects on plasma membrane curvature and by a direct interaction between lipid and protein. The GPCR neurotransmitter serotonin1A receptor has ten closely bound cholesterol molecules, and these control its organization and positioning; the receptor senses membrane cholesterol via a lysine residue in a so-called 'CRAC' motif in transmembrane helix 2. Ion pumps such as the (Na+-K+)-ATPase, which have specific binding sites for cholesterol molecules, are the single most important consumer of ATP in cells and are responsible for the ion gradients across membranes that are essential for many cellular functions; depletion of cholesterol in the plasma membrane deactivates these ion pumps. In the brain in addition to being essential for the structure of the myelin sheath, cholesterol is a major component of synaptic vesicles and controls their shape and functional properties. In the nucleus of cells, cholesterol is intimately involved in chromatin structure and function. The role of cholesterol together with sphingolipids in the formation of the transient membrane nano-domains known as rafts (see the specific web page for detailed discussion), is of crucial importance for the function of cells, while the interaction of cholesterol with ceramides is essential for the barrier function of the skin. Cholesterol Biosynthesis Cholesterol biosynthesis involves a highly complex series of at least thirty different enzymatic reactions, which were unraveled in large measure by Konrad Bloch and Fyodor Lynen, who received the Nobel Prize for their work on the topic in 1964. When the various regulatory, transport, and genetic studies of more recent years are taken into account, it is obvious that this is a subject that cannot be treated in depth here. The bare bones of mechanistic aspects are therefore delineated, which with the references listed below should serve as a guide to further study. In plants, cholesterol synthesis occurs by a somewhat different pathway with cycloartenol as the key intermediate. We'll explore the reaction mechanisms for several of the enzymes on this complicated pathway given its medical importance. Almost all nucleated cells are able to synthesize their full complement of cholesterol. The first steps involve the synthesis of the important intermediate mevalonic acid from acetyl-CoA and acetoacetyl-CoA, both of which are in fact derived from acetate, in two enzymatic steps. These precursors are in the cytosol as is the first enzyme, 3-hydroxy-3-methyl-glutaryl(HMG)-CoA synthase. The second enzyme HMG-CoA reductase is a particularly important control point, and is widely regarded as the rate-limiting step in the overall synthesis of sterols; its activity is regulated at the transcriptional level and by many more factors including a cycle of phosphorylation-dephosphorylation. This and subsequent enzymes are membrane-bound and are located in the endoplasmic reticulum. The enzyme HMG-CoA reductase is among the targets inhibited by the drugs known as ‘statins’ so that patients must then obtain much of their cholesterol from the diet. The first two reactions in the synthesis of cholesterol are shown in Figure \(5\). HMG-CoA Synthase We saw this reaction in the synthesis of ketone bodies in Chapter 17. This enzyme catalyzes the condensation of acetoacetyl–CoA (AcAc–CoA) and acetyl–CoA (Ac-CoA) to form 3-hydroxy-3-methylglutaryl (HMG)–CoA, and requires the formation of a C-C bond. HMG-CoA synthase forms a C-C bond by activating the methyl group of acetyl-cysteine. The acetyl group comes from an acyl-CoA "donor". The enzyme catalyzes the first committed step in the formation of complex isoprenoids (like cholesterol) and ketone bodies. The product, 3-hydroxy-3-methylglutaryl HMG–CoA, can either be reduced by HMG-CoA reductase to form mevalonate, which leads to cholesterol synthesis, or cleaved by the enzyme HMG-CoA lyase, to produce acetoacetate, a ketone body. HMG-CoA synthase catalyzes a bisubstrate reaction that displays ping-pong kinetics, characteristic of a covalent enzyme intermediate. The first substrate binds to the enzyme and transfers an acetyl group to a nucleophilic Cys 111 in the active site to form an acetyl-Cys 111 intermediate. Free CoA departs. Next the second substrate, acetoacetyl-CoA binds, and condenses with the acetyl group donated by acetyl-Cys 111. This condensation involves an enolate. A plausible reaction mechanism is shown in Figure \(6\). Figure \(6\): Reaction mechanism for HMG-CoA synthase Hence there are three parts of the reaction: acetylation/deacetylation, condensation/cleavage with an enolate intermediate, and C-C formation and hydrolysis/dehydration. Figure \(7\) shows an interactive iCn3D model of the Staphylococcus aureus HMG-COA Synthase with bound HMG-CoA and acetoacetyl-CoA (1XPK) Figure \(7\): Staphylococcus aureus HMG-COA Synthase with bound HMG-CoA and acetoacetyl-CoA (1XPK). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...deiFs7JceE5H76 The biologically active unit (homodimer) is shown with each monomer shown in a different color. The A chain (light cyan) has bound HMG-CoA (HMG) while the B chain (light gold) has acetoacetyl-CoA (CAA) bound. The Glu 79, Cys 111, and His 233 in each subunit are shown in CPK sticks and labeled. Note that the Cys 111 is covalently modified in each subunit. Figure \(8\) shows an interactive iCn3D model of the human 3-hydroxy-3-methylglutaryl CoA synthase I with bound CoASH (2P8U) Figure \(8\): Human 3-hydroxy-3-methylglutaryl CoA synthase I (monomer) with bound CoASH (2P8U). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...xxEoUfecTk3eL6 The active site side chains are numbered differently (Glu 95, Cys 129, and His 264) compared to the S. aurus protein. Only the monomer is shown in this model. HMG-CoA Reductase In this key rate-limiting step that commits HMG to the sterol and isoprenoid synthetic pathways, HMG is converted to mevalonate. A reducing agent (NADPH) is required for this biosynthetic reaction. A plausible mechanism is shown in Figure \(9\). Figure \(10\) shows an interactive iCn3D model of the catalytic domain of human HMG-CoA reductase with bound HMG-CoA (1DQ9). Figure \(10\): Catalytic domain of human HMG-CoA reductase with bound HMG-CoA (1DQ9). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...dZvWzTocKhUzt8 Two of the four identical monomers in the biological unit, the C (gray) and D (cyan) chains are shown. HMG-CoA is shown in spacefill and CPK colors. Three key catalytic residues, His 866 in the C chain (b chain in the mechanism above) and Lys 691 and Asp 767 in the D chain (A chain in the mechanism above) are shown in sticks, CPK colors and labeled. Synthesis of 5-isopentenyl and 2-isopentenyl pyrophosphate The next sequence of reactions involves first the phosphorylation of mevalonic acid by a mevalonate kinase to form the 5‑monophosphate ester, followed by further phosphorylation to yield an unstable pyrophosphate, which is rapidly decarboxylated to produce 5-isopentenyl pyrophosphoric acid, the universal isoprene unit. An isomerase converts part of the latter to 3,3-dimethylallyl pyrophosphoric acid. These reactions are shown in Figure \(11\). Two phosphorylations are required, one by mevalonate kinase, which proceeds by an ordered sequential binding of mevalonate as the first reactant and its phosphomevalonate as the first product released. The enzyme is inhibited by two downstream products of the reaction pathway, farnesyl pyrophosphate, and geranyl pyrophosphate. A mechanism for mevalonate kinase is shown in Figure \(12\). A reaction mechanism for the second kinase, phosphomevalonate kinase, showing progression through the transition state, is shown in Figure \(13\). The reaction proceeds through direct phosphorylation through a dissociative mechanism. Mevalonate diphosphate decarboxylase This enzyme catalyzes the decarboxylation of 5-pyrophosphate mevalonate to 5-isopentenylpyrophosphate as shown in Figure \(14\). The binding of the two substrates, pyrophosphatemevalonate (or mevalonate pyrophosphate - MVAPP) and ATP to the enzyme (A), and a reaction mechanism (B, are shown in Figure \(15\). Panel (a) shows changes in the enzyme structure upon substrate binding. The apo-enzyme is shown on the left. The middle structure shows the enzyme after binding of the first substrate, MVAPP. A key loop (β10-α4) is shown as a cyan surface. The enzyme is then shown in an open conformation with the second substrate (ATP) also bound. An additional phosphate-biding loop is shown in magenta. At the far right, the enzyme is shown in a closed conformation after conformational changes in both loops which traps substrates in the active site. These changes enable catalysis. Product release follows. Panel (b) shows the dissociative phosphoryl transfer catalytic mechanism. At the top-left, D282 is shown interacting with the 3′-OH group of MVAPP (red). The top-right shows a dissociative phosphoryl (blue) transition state. In the bottom-left, the phosphate attaches to the 3′ oxygen (red) of MVAPP. The bottom-right shows the products after the dephosphorylation and decarboxylation to produce IPP, ADP, phosphate, and CO2. K187 from the β10-α4 loop and metal ions in the active site are involved in neutralizing the negatively charged environment and assists catalysis. Figure \(16\) shows an interactive iCn3D model of mevalonate diphosphate decarboxylase with mevalonate-5-diphosphate, AMPPCP and Magnesium (6E2U). Figure \(16\): Mevalonate diphosphate decarboxylase with mevalonate-5-diphosphate, AMPPCP and Magnesium (6E2U) . (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...SukmP1BTyNhH26 The correctly positioned substrates interact with two Mg 2+ ions in the initial steps of the reaction. The conserved lysine facilitates the phosphoryl transfer. Polymerization of isoprene Isoprene, a small branched alkadiene, which can polymerize into larger molecules containing isoprene monomer to form isoprenoids, often called terpenes. Instead of using isoprene as the polymerization monomer, either dimethylally pyrophosphate (DMAPP) or isopentenylpyrophosphate (IPP) are used biologically. Figure \(\PageIndex{x}\) below shows how DMAPP and IPP (both containing 5Cs) are used in a polymerization reaction to form geranyl-pyrophosphate (C10), farnesyl pyrophosphate (C15) and geranyl-geranyl pyrophosphate (C20). Figure \(17\): The condensation of IPP and DMAPP is a head-to-tail condensation reaction. Another IPP reacts with geranylpyrophosphate using the same enzyme to produce farnesylpyrophosphate. DMAPP is first formed by the isomerization of an IPP to DMAPP catalyzed by isopentenyl-diphosphate delta-isomerase. It catalyzes the 1,3-allylic rearrangement of the homoallylic substrate isopentenyl (IPP). 5-isopentenyl pyrophosphate is a nucleophile, but its isomer, DMAPP, is highly electrophilic, which promotes the condensation of the two molecules. A mechanism for the next reaction, the first condensation of DMAPP and IPP to form geranylpyrophosphate by farnesyl pyrophosphate (diphosphate) synthase reaction, is shown in Figure \(18\). The reaction appears to proceed using a carbocation transition state, followed by the transfer of a hydrogen atom (not ion) from IPP to pyrophosphate. Figure \(19\) shows an interactive iCn3D model of E. Coli farnesyl pyrophosphate synthase bound to isopentyl pyrophosphate and dimethylallyl S-thiolodiphosphate (1RQI). Figure \(19\): Farnesyl pyrophosphate synthase Bound to isopentyl pyrophosphate (IPP) and dimethylallyl S-thiolodiphosphate (1RQI). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...kQBU5dNnKrCVt6 DST in the structure is dimethylallyl S-thiolodiphosphate, an analog of dimethylallyl diphosphate (DMAPP). Key conserved amino acids involved in substrate binding, transition state stabilization, and catalysis are shown as sticks, CPK colors. Thr203, Gln241, and Lys202 presumably stabilized the carbocation intermediate/transition state. Lys202 also forms a hydrogen bond with DMAPP. Two arginines (69, 116) form salt bridges with the pyrophosphates of IPP and DMAPP. Many isoprenoid lipids are made from farnesyl pyrophosphate. For membrane purposes, the most important of these is cholesterol. Figure \(20\) shows an overview of the synthesis of cholesterol from two farnesyl pyrophosphates linking together in a "tail-to-tail" reaction to form squalene, a precursor of cholesterol. Each isoprene unit (5Cs) is shown in different colors to make it easier to see. Figure \(20\): Synthesis of squalene from isoprene units Figure \(21\) shows reactants (two farnesyl pyrophosphate), intermediate (presqualene diphosphate) and product (squalene) in the reaction catalyzed by squalene synthase. In the squalene synthase reaction, two molecules of farnesyl pyrophosphate condense to yield presqualene pyrophosphate. In turn, this is reduced by NADPH to produce the key intermediate squalene. The enzyme squalene synthase, which regulates the flow of metabolites into either the sterol or non-sterol pathways (with farnesyl pyrophosphate as the branch point), is considered to be the first committed enzyme in cholesterol biosynthesis. Given the importance of this reaction, we will explore the unique mechanism of squalene synthase in some detail. Part 1: Formation of the cyclopropyl presqualene intermediate Figure \(22\) shows the mechanism for the first part of the reaction in which the cyclopropyl intermediate presqualene form. The reaction proceeds through a series of carbocation intermediates. Part 2: Conversion of cyclopropyl intermediate to squalene - reduction The next part of the reaction involves the reductive formation of squalene, as shown in Figure \(23\). Figure \(24\) shows an interactive iCn3D model of human squalene synthase with bound inhibitor (1EZF). Figure \(24\): Human squalene synthase with bound inhibitor (1EZF). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...jLreHkvbftoFPA Tyr 171 acts as a general acid/base in the first half of the reaction. Arg 218 and 228 stabilize the diphosphate in the transition state as it leaves. Phe 288 stabilizes the reactive carbocation. The enzyme has a single domain that contains a large channel in one face of the enzyme which leads from a solvent-exposed to the hydrophobic interior. Two FPPs bind in the beginning of the channel where key side chains for the first reaction are located. The cyclopropyl intermediate then moves into the hydrophobic end of the channel where it reacts in the second half of the reaction without exposure to water. In the next important step, squalene is oxidized by a squalene monooxygenase to squalene 2,3-epoxide, a key control point in the cholesterol synthesis pathway. This introduces the oxygen atom to squalene that becomes the signature oxygen of the hydroxyl group in cholesterol. The epoxide then undergoes cyclization catalyzed by the enzyme squalene epoxide lanosterol-cyclase to form the first steroidal intermediate lanosterol (or cycloartenol en route to phytosterols in photosynthetic organisms). This is illustrated in Figure \(26\). In this remarkable reaction, there is a series of concerted 1,2-methyl group and hydride shifts along the chain of the squalene molecule to bring about the formation of the four rings. No intermediate compounds have been found. This is believed to be one of the most complex single enzymatic reactions ever to have been identified, although the enzyme involved is only 90 kDa in size. Again, the reaction takes place in the endoplasmic reticulum, but a cytosolic protein, sterol carrier protein 1, is required to bind squalene in an appropriate orientation in the presence of the cofactors NADPH, flavin adenine dinucleotide (FAD) and O2; the reaction is promoted by the presence of phosphatidylserine. The ring closure reaction starting with the epoxide involves a concerted flow of electrons from a source to the epoxide oxygen atom electron sink. This brings to mind a reaction known to all students who have ever studied chemistry, the reaction of the pH indicator phenolphthalein with a base to produce a pink colored-solution. That reaction which produces a more conjugated molecule that absorbs at 553 nm (green) which causes a magenta solution color, is illustrated for comparison (and fun) in Figure \(27\). In subsequent steps, lanosterol is converted to cholesterol by a series of demethylations, desaturations, isomerizations, and reductions, involving nineteen separate reactions as illustrated in Figure \(28\). Thus, demethylation reactions produce zymosterol as an intermediate, and this is converted to cholesterol via a series of intermediates, all of which have been characterized, and by at least two pathways that utilize essentially the same enzymatic machinery but differ in the order of the various reactions, mainly at the point at which the Δ24 double bond is reduced. Desmosterol is the key intermediate in the so-called 'Bloch' pathway, while 7‑dehydrocholesterol is the immediate precursor in the 'Kandutsch-Russell' pathway. While some tissues, such as adrenal glands and testis, use the Bloch pathway mainly, the brain synthesizes much of its cholesterol by the 'Kandutsch-Russell' pathway. This may enable the production of a variety of other minor sterols for specific biological purposes in different cell types/locations. The energy cost of the synthesis of one cholesterol molecule is roughly one hundred ATP equivalents, and eleven oxygen molecules are required. Synthesis occurs mainly in the liver, although the brain (see below), peripheral nervous system, and skin synthesize their own considerable supplies. Cholesterol is exported from the liver and transported to other tissues in the form of low-density lipoproteins (LDL) for uptake via specific receptors. In animals, cells can obtain the cholesterol they require either from the diet via the circulating LDL, or they can synthesize it themselves as outlined above. Cholesterol biosynthesis is highly regulated with rates of synthesis varying over hundreds of fold depending on the availability of any external sources of cholesterol, and cholesterol homeostasis requires the actions of a complex web of enzymes, transport proteins, and membrane-bound transcription factors, as discussed below. Regulation of Cholesterol Homeostasis In humans, only about a third of the cholesterol is of dietary origin (mainly eggs and red meat), the remainder is produced by synthesis de novo in the endoplasmic reticulum. The latter must be tightly regulated as it is an energetically expensive process that requires appreciable amounts of acetyl-CoA, ATP, oxygen, and the reducing factors NADPH and NADH, especially since cholesterol cannot be catabolized for energy purposes (see below). Many factors are involved in maintaining the large differences in cholesterol concentrations among the various membranes and organelles in cells within precise limits. In order to explain how cholesterol in the plasma membrane, where it is most abundant, can regulate cholesterol biosynthesis and uptake through enzymes in the endoplasmic reticulum, where it is least abundant, it has been suggested that a key to the process is that there are three pools of cholesterol in the plasma membrane with distinct functional roles. The first of these is “accessible” to receptor proteins for transport to the endoplasmic reticulum, while the second pool is sequestered by sphingomyelin and can be released by the action of sphingomyelinase if required. The third residual pool of cholesterol is essential for plasma membrane integrity. These correspond to about 16, 15, and 12 mol % of total plasma membrane lipids, respectively, in cholesterol-replete cells. Simplistically, when cholesterol in the plasma membrane is in excess for any reason, e.g., after LDL uptake by receptor-mediated endocytosis, there is a rise in accessible cholesterol, which is then transported to the endoplasmic reticulum to switch off cholesterol biosynthesis and expression of the LDL receptor. This process requires a host of regulatory proteins and mechanisms that can involve either vesicle formation or non-vesicular pathways that utilize specific transport proteins, such as the ABC transporters. Ultimately, post-translational control of the many different enzymes involved provides a rapid means for modifying flux through the biosynthetic pathway in the endoplasmic reticulum; some are rapidly degraded in response to tissue levels of cholesterol and its intermediates, while others have their activity altered through phosphorylation or acetylation mechanisms. For example, the second rate-limiting enzyme in cholesterol biosynthesis is squalene monooxygenase, which undergoes cholesterol-dependent proteasomal degradation when cholesterol is in excess, guided by a 12-amino acid hydrophobic sequence on the enzyme that can serve as a degradation signal. When the cholesterol concentration in the endoplasmic reticulum is high, the degradation sequence detaches from the membrane and is exposed to provide the signal for the enzyme to be degraded. Similarly, HMG-CoA reductase is recognized as the key enzyme in the regulation of cholesterol biosynthesis, and this can be regulated by a feedback mechanism involving ubiquitin–proteasomal degradation. Further regulation of cholesterol biosynthesis is exerted by sterol intermediates in cholesterol biosynthesis, such as lanosterol and 24,25‑dehydrolanosterol (dimethyl-sterols) by accelerating degradation of the biosynthetic enzymes such as HMG-CoA reductase. It is noteworthy that ceramide down-regulates cholesterol synthesis – another link between cholesterol and sphingolipid metabolism. The regulatory element-binding proteins (mainly SREBP-1c and SREBP-2), which contain an N-terminal membrane domain and a C-terminal regulatory domain, are essential to the maintenance of cholesterol levels. Each is synthesized as an inactive precursor that is inserted into the endoplasmic reticulum where it can encounter an escort protein termed SREBP cleavage-activating protein (SCAP), which is the cellular cholesterol sensor. When the latter recognizes that cellular cholesterol levels are inadequate, it binds to the regulatory domain of SREBP. The SCAP-SREBP complex then moves to the Golgi, where two specific proteases (designated site-1 and site-2 proteases) cleave the SREBP enabling the C-terminal regulatory domain to enter the nucleus. There it activates transcription factors, such as the nuclear liver X receptor (LXR), which stimulate the expression of the genes coding for the LDL receptor in the plasma membrane and for the key enzyme in cholesterol biosynthesis, HMG-CoA reductase. This in turn stimulates the rate of cholesterol uptake and synthesis. Conversely, when cholesterol in the endoplasmic reticulum exceeds a threshold, it binds to SCAP in such a way that it prevents the SCAP-SREBP complex from leaving the membrane for the nucleus, cholesterol synthesis and uptake are thereby repressed, and cholesterol homeostasis is restored. In effect, cholesterol exerts feedback inhibition by suppressing its own production by preventing the proteolytic cleavage and maturation of SREBP-2. Oxysterols, especially 25-hydroxycholesterol, are also inhibitors of this process. Cholesterol in the endoplasmic reticulum is transferred to the Golgi and eventually to the plasma membrane by vesicular and non-vesicular transport mechanisms involving in part soluble sterol transport proteins, including the so-called 'START' domain proteins, and partly by binding to those proteins that are intimately involved in the transport and metabolism of polyphosphoinositides such as phosphatidylinositol 4-phosphate (PI(4)P). In the latter mechanism, cholesterol is transported by binding to the ORD domain of oxysterol binding protein (OSBP) or Osh4 in yeast, before OSBP binds to PI(4)P in the plasma membrane to transfer its cargo. The key to this process is that cholesterol and PI(4)P are synthesized at two different locations, i.e., the endoplasmic reticulum for sterols and the trans-Golgi network and plasma membrane for PI(4)P, so the two lipids do not compete but rather can be exchanged. OSBP carries cholesterol in the forward direction to the trans-Golgi network and plasma membrane and PI(4)P, which binds to a C-terminal PH domain in the protein, in the reverse direction. The subsequent hydrolysis of PI(4)P is the energy source for the reaction, and indeed PI(4)P has been termed "lipid ATP". As this reaction is irreversible, a gradient of cholesterol along organelles of the secretory pathway is established. OSBP is thus a lipid transfer protein that enables two organelles to exchange cholesterol rapidly between them at membrane contact sites in a cycle of reactions involving membrane tethering, cholesterol transport, PI(4)P counter transport, and PI(4)P hydrolysis. A similar mechanism is involved in the transport of phosphatidylserine from the endoplasmic reticulum to the inner leaflet of the plasma membrane. Subsequently, the ATP binding cassette (ABC) transporters ABCA1 and ABCG1 in the plasma membrane, which contains much of the cellular cholesterol, are activated to export the excess. Nuclear factor erythroid 2 related factor-1 or NRF1 in the endoplasmic reticulum binds directly to cholesterol and senses when its level is high to bring about a de-repression of genes involved in cholesterol removal, also with mediation by the liver X receptor. Also, side-chain oxysterols, especially 25-hydroxycholesterol, can suppress the activation of SREBP by binding to an oxysterol-sensing protein in the endoplasmic reticulum. Within cells, cholesterol derived initially from the lysosomal degradation of low-density lipoproteins is transferred first to the plasma membrane and thence to the endoplasmic reticulum, the latter step by a mechanism involving proteins known as GRAMD1s embedded in the endoplasmic reticulum membrane at sites in contact with the plasma membrane. These have two functional domains: the START-like domain that binds cholesterol and the GRAM domain that binds anionic lipids, such as phosphatidylserine, and so are able to form a link between the two membranes that enable the transfer of cholesterol. In peripheral tissues, excess cholesterol is exported to high-density lipoproteins (HDL) in the circulation and returned to the liver, a process known as reverse cholesterol transport. The liver is important for cholesterol synthesis, but it is essential for its elimination from the body in bile. Also, some lipoproteins with their content of cholesterol and cholesterol esters are delivered to lysosomes by endocytosis for degradation. The cholesterol is transported to the inner surface of the lysosomal membrane through the glycocalyx, via a transglycocalyx tunnel, with the aid of Niemann-Pick C1, C2, and other proteins, and thence via contact sites between membranes to other organelles. Cholesterol in cellular membranes in excess of the stoichiometric requirement can escape back into the cell, where it may serve as a feedback signal to down-regulate cholesterol accumulation, while some is converted to the relatively inert storage form, i.e., cholesterol esters, and some is used for steroidogenesis. The intestines play a major part in cholesterol homeostasis via the absorption of dietary cholesterol and fecal excretion of cholesterol and its metabolites. A specific transporter (Niemann-Pick C1-like 1 or NPC1L1)in the brush border membrane of enterocytes in the proximal jejunum of the small intestine is involved in the uptake of cholesterol from the intestinal contents, while the metabolism of sterols in the intestines is controlled mainly by an acetyl-CoA acetyltransferase (ACAT2), which facilitates intracellular cholesterol esterification, and the microsomal triglyceride transfer protein (MTTP), which is involved in the assembly of chylomicrons for export into lymph. Some cholesterol can be transferred in the opposite direction (trans-intestinal cholesterol excretion), but the quantitative importance of this process is not clear. There is evidence that dietary or synthesized cholesterol is necessary to maintain intestinal integrity, as cholesterol derived from circulating lipoproteins is not sufficient for the purpose. In the intestines and especially the colon, the intestinal microflora are able to hydrogenate cholesterol from bile, diet, and desquamated cells to form coprostanol with an efficiency that is dependent on the composition of microbial species. Coprastanol is not absorbed by the intestinal tissue to a significant extent, and it may inhibit the uptake of residual cholesterol. There are two mechanisms for this conversion in bacteria, one involving direct reduction and another via cholestenone and coprostanone as intermediates, and as the relevant genes have now been identified the therapeutic potential is under investigation. Brain: There are substantial differences in cholesterol synthesis and metabolism in brain in comparison to the liver and peripheral tissues. Trace amounts only of cholesterol are able to cross the blood-brain barrier via transport in low-density lipoproteins. Therefore, virtually all the cholesterol in the brain must be synthesized de novo, mainly in astrocytes (glial cells). During the perinatal and adolescent years especially, cholesterol is synthesized in large amounts to form the myelin that surrounds the axons, before this rate begins to decline to eventually reach about 10% of earlier values. Cholesterol is transported to neurons in the form of Apo E complexes in discoidal HDL-like particles, for which seven main receptors have been identified in brain cells that take up cholesterol from these lipoproteins. Apo E is synthesized in the brain, and its transcription is regulated by 24-hydroxy-cholesterol concentrations. Similarly, in the brain and central nervous system, cholesterol synthesis is regulated independently of that in peripheral tissues, mainly by forms of the liver X receptor (LXR). As cholesterol and oxysterols are involved in providing neuroprotective effects and lowering neuroinflammation, dysregulation of their concentrations has been noted in many neurodegenerative disorders. Most of the lipoproteins in cerebrospinal fluid differ from the nascent poorly-lipidated HDL secreted by astrocytes, suggesting that the latter are modified during maturation. Cholesterol Catabolism Cholesterol is not readily degraded in animal tissues so does not serve as a metabolic fuel to generate ATP. Only the liver possesses the enzymes to degrade significant amounts, and then via pathways that do not lead to energy production. Cholesterol and oxidized metabolites (oxysterols) are transferred back from peripheral tissues in lipoprotein complexes to the liver for catabolism by conversion to oxysterols and bile acids. The latter are exported into the intestines to aid in digestion, while leading to some loss that is essential for cholesterol homeostasis. Until recently, it was believed that approximately 90% of cholesterol elimination from the body occurred via bile acids in humans. However, experiments with animal models now suggest that a significant amount is secreted directly into the intestines by a process known as trans-intestinal cholesterol efflux. How this occurs and its relevance to humans are under active investigation. Gut bacteria reduce some of the cholesterol in the diet to highly insoluble 5β-cholestan-3β-ol (coprostanol), which is excreted and can be used as a biomarker for sewage in the environment. Certain bacterial species contain a 3β-hydroxysteroid:oxygen oxidoreductase (EC 1.1.3.6), commonly termed cholesterol oxidase, a flavoenzyme that catalyzes the oxidation of cholesterol to cholest-5-en-3-one which is then rapidly isomerized to cholest-4-en-3-one as the first essential step in the catabolism of sterols. The enzyme is widespread in organisms that degrade organic wastes, but it is also present in pathogenic organisms where it influences the virulence of infections (see below). In biotechnology, it has been used for the production of a number of steroids, and it is employed in a clinical procedure for the determination of cholesterol levels in serum. Cholesterol Esters Cholesterol esters, i.e., with long-chain fatty acids linked to the hydroxyl group, are much less polar than free cholesterol and appear to be the preferred form for transport in plasma and as a biologically inert storage or detoxification form to buffer an excess. They do not contribute to membrane structures but are packed into intracellular lipid droplets. Cholesterol esters are major constituents of the adrenal glands, and they accumulate in the fatty lesions of atherosclerotic plaques. Similarly, esters of steroidal hormones are also present in the adrenal glands, where they are concentrated in cytosolic lipid droplets adjacent to the endoplasmic reticulum; 17β-estradiol, the principal estrogen in fertile women, is transported in lipoproteins in the form of a fatty acid ester. Because of the mechanism of synthesis (see below), plasma cholesterol esters tend to contain relatively high proportions of the polyunsaturated components typical of phosphatidylcholine as shown in Table \(1\) below. Arachidonic and “adrenic” (20:4(n-6)) acids can be especially abundant in cholesterol esters from the adrenal gland. Table \(1\). Fatty acid composition of cholesterol esters (wt % of the total) from various tissues. Form Fatty acids 16:0 18:0 18:1 18:2 18:3 20:4 22:4 Human plasma 12 2 27 45   8 liver 23 10 28 22   6 Sheep plasma 10 2 27 35 7 - - liver 17 9 29 7 4 3 - adrenals 13 7 35 18 2 4 2 Data from - Christie, W.W. et al. Lipids, 10, 649-651 (1975); DOI. Nelson, G.J. Comp. Biochem. Physiol., 30, 715-725 (1969); Horgan, D.J. and Masters, C.J. Aust. J. Biol. Sci., 16, 905-915 (1963); Nestel, P.J. and Couzens, E.A. J. Clin. Invest., 45, 1234-1240 (1966); DOI. In plasma and in the high-density lipoproteins (HDL) in particular, cholesterol esters are synthesized largely by the transfer of fatty acids to cholesterol from position sn-2 of phosphatidylcholine (‘lecithin’) catalyzed by the enzyme lecithin:cholesterol acyl transferase (LCAT); the other product is 1-acyl lysophosphatidylcholine. This is illustrated in Figure \(29\). In fact, the reaction occurs in several steps. First, apoprotein A1 in the HDL acts to concentrate the lipid substrates near LCAT and present it in the optimal conformation; at the same time, it opens a lid on the enzyme that activates it by opening up the site of transesterification. Then, cleavage of the sn-2 ester bond of phosphatidylcholine occurs via the phospholipase activity of LCAT with the release of a fatty acyl moiety. This is transacylated to the sulfur atom of a cysteine residue forming a thioester, and ultimately it is donated to the 3β-hydroxyl group of cholesterol to form the cholesterol ester. Some LCAT activity has also been detected in apolipoprotein B100-containing particles (β-LCAT activity as opposed to α-LCAT with HDL). It has been established that human LCAT is a relatively small glycoprotein with a polypeptide mass of 49 kDa, increased to about 60 kDa by four N-glycosylation and two O-glycosylation moieties. Most of the enzyme is produced in the liver and circulates in the bloodstream bound reversibly to HDL, where it is activated by the main protein component of HDL, apolipoprotein A1. As cholesterol esters accumulate in the lipoprotein core, cholesterol is removed from its surface thus promoting the flow of cholesterol from cell membranes into HDL. This in turn leads to morphological changes in HDL, which grow and become spherical. Subsequently, cholesterol esters are transferred to the other lipoprotein fractions LDL and VLDL, a reaction catalyzed by cholesterol ester transfer protein. This process promotes the efflux of cholesterol from peripheral tissues (‘reverse cholesterol transport’), especially from macrophages in the arterial wall, for subsequent delivery to the liver. LCAT is often stated to be the main driving force behind this process, and it is of great importance for cholesterol homeostasis and a suggested target for therapeutic intervention against cardiovascular disease. The stereospecificity of LCAT changes with molecular species of phosphatidylcholine containing arachidonic or docosahexaenoic acid, when 2-acyl lysophosphatidylcholines are formed. This reaction may be especially important for the supply of these essential fatty acids to the brain in that such lysophospholipids are believed to cross the blood-brain barrier more readily than the free acids. In other animal tissues, a further enzyme acyl-CoA:cholesterol acyltransferase (ACAT) synthesizes cholesterol esters from CoA esters of fatty acids and cholesterol. ACAT exists in two forms, both of which are intracellular enzymes found in the endoplasmic reticulum and are characterized by multiple transmembrane domains and a catalytic histidine residue in a hydrophobic domain; they are members of the O-acyltransferase (MBOAT) superfamily. ACAT1 is present in many tissues, but especially in macrophages and adrenal and sebaceous glands, which store cholesterol esters in the form of cytoplasmic lipid droplets; it is responsible for the synthesis of cholesterol esters in arterial foam cells in human atherosclerotic lesions. ACAT2 is found only in the liver and small intestine, and it is believed to be involved in the supply of cholesterol esters to the nascent lipoproteins. Analogous enzymes are found in yeast where ergosterol is the main sterol, but a very different process occurs in plants. Oxidized Cholesterol Esters: All lipid classes containing polyunsaturated fatty acids are susceptible to oxidation. Under normal circumstances, cholesterol esters are considered to be relatively inert. However, when they contain oxidized polyunsaturated fatty acids, their properties change and they acquire biological activity. Such oxidized cholesterol esters may be formed by a reaction with 15‑lipoxygenase, but they can be produced also through free radical-induced lipid peroxidation, and they have been detected in lipoproteins, LDL especially, in human blood and atherosclerotic lesions. Those oxidized cholesterol esters in plasma are trafficked into cells and metabolized by the same mechanisms as the corresponding unoxidized lipids. Such "minimally oxidized LDL" do not bind to CD36 but rather to CD14, a receptor that recognizes bacterial lipopolysaccharides. The result is stimulation of toll-like receptor 4 (TLR4), although the response differs from that of lipopolysaccharides. In addition, oxidized metabolites of cholesteryl arachidonate of this kind stimulate macrophages to express inflammatory cytokines of relevance to atherosclerosis among other effects. Oxidized cholesterol esters can be hydrolyzed to release their fatty acids, which can then be incorporated into phospholipids with a different repertoire of activities. Hydrolysis of cholesterol esters: Cholesterol ester hydrolases in animals liberate cholesterol and free fatty acids when required for membrane and lipoprotein formation, and they also provide cholesterol for hormone synthesis in adrenal cells. Many cholesterol ester hydrolases have been identified, including a carboxyl ester hydrolase, a lysosomal acid cholesterol ester lipase, hormone-sensitive lipase, and hepatic cytosolic cholesterol ester hydrolase. These are located in many different tissues and organelles and have multiple functions. A neutral cholesterol ester hydrolase has received special study, as it is involved in the removal of cholesterol esters from macrophages so reducing the formation of foam cells and thence the development of fatty streaks within the arterial wall, a key event in the progression of atherosclerosis. Other Animal Sterols Cholesterol will oxidize slowly in tissues or foods to form a range of different products with additional hydroperoxy, epoxy, hydroxy or keto groups, and these can enter tissues via the diet. There is increasing interest in these from the standpoint of human health and nutrition since the accumulation of oxo-sterols in plasma is associated with inhibition of the biosynthesis of cholesterol and bile acids and with other abnormalities in plasma lipid metabolism. A number of other sterols occur in small amounts in tissues, most of which are intermediates in the pathway from lanosterol to cholesterol, although some of them have distinct functions in their own right. Lanosterol, the first sterol intermediate in the biosynthesis of cholesterol, was first found in wool wax, both in free and esterified form, and this is still the main commercial source. It is found at low levels only in most other animal tissues (typically 0.1% of the cholesterol concentration). As oxygen is required, lanosterol cannot be produced by primitive organisms, hence its absence from prokaryotes, leading to some speculation on its evolutionary significance. When sterols became available to eukaryotes, much greater possibilities opened for their continuing evolution. The production of cholesterol from lanosterol is then seen as ‘molecular streamlining’ by evolution, removing protruding methyl groups that hinder the interaction between sterols and phospholipids in membranes. Desmosterol (5,24-cholestadien-3β-ol), the last intermediate in the biosynthesis of cholesterol by the Bloch pathway, may be involved in the process of myelination, as it is found in relative abundance in the brains of young animals but not in those of adults, other than astrocytes. It is also found in appreciable amounts in testes and spermatozoa together with another cholesterol intermediate, testis meiosis-activating sterol. In addition, there is evidence that desmosterol activates certain genes involved in lipid biosynthesis in macrophages, and may deactivate others associated with the inflammatory response. There is a rare genetic disorder in which there is an impairment in the conversion of desmosterol to cholesterol, desmosterolosis, with serious consequences in terms of mental capacity. These and related sterols appear to be essential for human reproduction. In human serum, the levels of lathosterol (5α-cholest-7-en-3β-ol) were found to be inversely related to the size of the bile acid pool, and in general, the concentration of serum lathosterol is strongly correlated with the cholesterol balance under most dietary conditions. The isomeric saturated sterols, cholestanol, and coprastanol, which differ in the stereochemistry of the hydrogen atom on carbon 5, are formed by microbial biohydrogenation of cholesterol in the intestines, and together with cholesterol are the main sterols in feces. Further examples of animal sterols include 7-dehydrocholesterol (cholesta-5,7-dien-3β-ol) in the skin, which on irradiation with UV light is converted to vitamin D3 (cholecalciferol). These sterols are shown in Figure \(30\). Figure \(30\): Other animal sterols Marine invertebrates produce a large number of novel sterols, with both unusual nuclei and unconventional sidechains, some derived from cholesterol and others from plant sterols or alternative biosynthetic intermediates. For example, at least 80 distinct sterols have been isolated from echinoderms and 100 from sponges. Cholesterol and Disease Elevated cholesterol and cholesterol ester levels are associated with the pathogenesis of cardiovascular disease (atherosclerotic plaques, myocardial infarctions, and strokes), as is well known, and this is considered briefly on this website together with the metabolism of the plasma lipoproteins. The rate-limiting enzyme in the synthesis of cholesterol HMG-CoA reductase is the target of statins, but drugs that target other steps in the biosynthetic pathway, especially the squalene monooxygenase and lanosterol synthase, are under investigation. Further discussion of such a complex nutritional and clinical topic is best left to others better qualified than myself. Cholelithiasis or the presence of 'stones' in the gallbladder or bile ducts, which consist largely of cholesterol (~85%), is one of the most prevalent and costly digestive diseases in developed countries. The primary cause is the excessive excretion of cholesterol from the liver. Excess accumulation of cholesterol associated with the metabolism of bis(monoacylglycero)phosphate and causing disturbances in glycosphingolipid trafficking in cell membranes is involved in the pathogenesis of Niemann-Pick C disease, a lysosomal storage disease in which endocytosed cholesterol becomes sequestered in late endosomes/lysosomes because of gene mutations affecting two binding proteins (NPC1 and NPC2) thereby causing neuronal and visceral atrophy. In addition, deficiencies in cholesterol transport and metabolism are associated with many forms of neurodegeneration, including Alzheimer’s disease, Huntington’s disease, and related conditions associated with old age. These proteins are also key virulence factors for several viral and bacterial pathogens. Several genetic disorders of cholesterol biosynthesis have been identified in recent years that can result in developmental malformations including neurologic defects. As there is limited cholesterol transport across the placenta, the human fetus is highly dependent upon endogenous synthesis. While the molecular basis for the altered developmental pathways is not fully understood, impaired synthesis of the hedgehog family of signaling proteins, which require covalently linked cholesterol to function in membranes, is believed to be involved in many cases. In others, there are confirmed enzyme defects. For example, the recessive Smith-Lemli-Opitz syndrome in infants born with a decreased concentration of the enzyme 7-dihydrocholesterol reductase, produces symptoms varying from mild autism to severe mental and often fatal physical problems. The effects are due to a lack of cholesterol and the accumulation of 7-dehydrocholesterol and its 27-hydroxy metabolite, as brain tissue cannot utilize dietary cholesterol or that produced peripherally. In fact, at least eight different inherited disorders of cholesterol biosynthesis lead to congenital abnormalities in those afflicted. In animal models, deficiencies in SREBP-2 and genes encoding sterol biosynthetic enzymes display embryonic lethality. Deficiencies in the enzymes responsible for the hydrolysis of cholesterol esters, such as the lysosomal acid lipase, occur in Wolman disease and cholesterol ester storage disease. Cholesterol and other sterols bind directly to several immune receptors, especially in macrophages and T cells, and dynamic changes in cholesterol biosynthesis impact directly upon innate and adaptive immune responses, such that functional coupling between sterol metabolism and immunity has implications for health and disease. For example, cholesterol binds directly to the αβ T cell antigen receptor (αβTCR) and has at least two opposing functions in its activation. By binding to the trans-membrane domain of this receptor, it is kept in an inactive, non-signaling conformation, but when required it can stimulate the formation of receptor nanoclusters to increase their avidity for the antigen. In cancer, there is a high demand for cholesterol in order to support the inherent nature of tumor cells to divide and proliferate, and perturbations of reverse cholesterol transport can have negative consequences. Drugs that lower cholesterol levels in cancer cells by inhibiting the mevalonate pathway are undergoing clinical trials. When increased levels of sterols other than cholesterol are found in plasma, they usually serve as markers for abnormalities in lipid metabolism associated with disease states. For example, premature atherosclerosis and xanthomatosis occur in two rare lipid storage diseases, cerebrotendinous xanthomatosis, and sitosterolemia. In the former, cholestanol is present in all tissues, while in the latter, the dietary plant sterols campesterol and sitosterol accumulate in plasma and red blood cells. Inhibition of cholesterol biosynthesis may be associated with the appearance of some of the precursor sterols in the plasma. In infections with Mycobacterium tuberculosis, the organism uses host cholesterol as the major carbon and energy source and thereby promotes persistent infection with appreciable effects on pathogenicity. Similarly, Chlamydia trachomatis, a gram-negative obligate intracellular bacterium and a major cause of sexually transmitted infections, requires host cholesterol for growth. Many viruses use cholesterol as part of their life cycle, and reduction in cellular cholesterol is sometimes seen as an anti-viral strategy, although this may not always be helpful. For example, an HIV protein has a binding site for cholesterol, which it utilizes to facilitate the fusion with raft regions in the membranes of the host cell. Sterols: 2. Oxysterols and Other Cholesterol Derivatives Oxysterols as defined and discussed here are oxygenated derivatives of cholesterol and its precursors, i.e., with additional hydroxyl, epoxyl, or keto groups, that are found in all animal tissues. Many of these have vital functions in animals, while others are important as short-lived intermediates or end products in the catabolism or excretion of cholesterol or in the biosynthesis of steroid hormones, bile acids, and 1,25‑dihydroxy-vitamin D3. They are normally present in biological membranes and lipoproteins at trace levels only, though they can exert profound biological effects at these concentrations. However, they are always accompanied by a great excess (as much as 106-fold) of cholesterol per se. A multiplicity of different oxysterols are synthesized in cells by sequential reactions with specific oxygenases. However, because of the presence of the double bond in the 5,6-position, oxysterols can also be formed rapidly by non-enzymatic oxidation (autoxidation) of cholesterol and cholesterol esters within tissues with the formation of many different oxygenated derivatives. Simplistically, non-enzymatic oxidation leads mainly to the generation of products in which the sterol ring system is oxidized, while enzymatic processes usually produce metabolites with an oxidized side chain (7-hydroxylation is an important exception). Oxidized cholesterol molecules can also be generated by the gut microflora and be taken up through the enterohepatic circulation. Once an oxygen function is introduced into cellular cholesterol, the product can act as a biologically active mediator by interacting with specific receptors before it is metabolized to bile acids or is degraded further, processes assisted by the fact that oxysterols are able to diffuse much more rapidly through membranes than is cholesterol itself. Cholesterol metabolites of this kind are especially important in the brain, which is a major site for cholesterol synthesis de novo, and they are crucial elements of cholesterol homeostasis. Enzymatic Oxidation of Cholesterol Within animal cells, the oxidation of sterols is mainly an enzymic process that is carried out by several enzymes that are primarily from the cytochrome P450 family of oxygenases (named for a characteristic absorption at 450 nm). These comprise a disparate group of proteins that contain a single heme group and have a similar structural fold, though the amino acid sequences can differ appreciably. They are all mono-oxygenases. Oxysterol biosynthesis can be considered in terms of different pathways that depend on the position of the initial oxidation, but these pathways tend to overlap and lead to a complex web of different oxysterols (and eventually to bile acid formation). As these enzymes, which include cytochrome P450, cholesterol hydroxylase, hydroxysteroid dehydrogenases, and squalene epoxidase, are specific to particular tissues and indeed animal species, there is considerable variation in oxysterol distributions between organs. A few examples only of the first steps in some of these pathways are illustrated in Figure \(31\). As an example, a primary product is 7α‑hydroxycholesterol, which is an important intermediate in the biosynthesis of bile acids by the 'neutral' pathway and of many other oxysterols, and it is produced in the liver by the action of cholesterol 7α-hydroxylase (CYP7A1), an enzyme that has a critical role in cholesterol homeostasis. The reaction is under strict regulatory control, and the expression of CYP7A1 is controlled by the farnesoid X receptor (FXR) and is activated by cholic and chenodeoxycholic acids. Any circulating 7α‑hydroxycholesterol represents leakage from the liver. Further oxidation of 7α‑hydroxycholesterol can occur, and the action of CYP3A4 in humans generates 7α,25‑dihydroxycholesterol as an important metabolite, for example, while oxidation by CYP27A1 yields 7α,27‑dihydroxycholesterol; the latter is regarded as a key step in a further pathway to oxysterols and bile acids. On the other hand, the epimer 7β‑hydroxycholesterol is produced in the brain by the action of the toxic β-amyloid peptide and its precursor on cholesterol, but whether this is involved in the pathology of Alzheimer’s disease has yet to be determined. The hydroxysteroid 11-β-dehydrogenase 1 (HSD11B1) is responsible for the conversion of 7β-hydroxy-cholesterol to the important metabolite 7-keto-cholesterol, while HS11B2 catalyzes the reverse reaction; 7-keto-cholesterol is also formed by autoxidation (see below). HSD11B1 is better known as the oxidoreductase that converts inactive cortisone to the active stress hormone cortisol in glucocorticoid target tissues. An alternative ('acidic') pathway to bile acids starts with the synthesis of 27-hydroxycholesterol (or more systematically named (25R)26‑hydroxycholesterol), which is produced by the cytochrome P450 enzyme (CYP27A1) and introduces the hydroxyl group into the terminal methyl carbon (C27 or C26 - used interchangeably). While this enzyme is present in the liver, it is found in many extra-hepatic tissues and especially the lung, which provides a steady flux of 27‑oxygenated metabolites to the liver. As a multifunctional mitochondrial P450 enzyme in the liver, it generates both 27‑hydroxycholesterol and 3β‑hydroxy-5-cholestenoic acid, the bile acid precursor, which occurs in small but significant amounts in plasma. 27‑Hydroxycholesterol is the most abundant circulating oxysterol, and its concentration in plasma correlates with that of total cholesterol. It can be oxidized to 7α,27‑dihydroxycholesterol by the enzyme CYP7B1. 4β‑Hydroxycholesterol is also abundant in plasma and is relatively stable; it is produced in humans by the action of the cytochromes CYP3A4 and CY3A5. In humans, the specific cytochrome P450 that produces 24S-hydroxycholesterol (cholest-5-ene-3β,24-diol) is cholesterol 24S‑hydroxylase (CYP46A1) and is located almost entirely in the smooth endoplasmic reticulum of neurons in the brain, including those of the hippocampus and cortex, which are important for learning and memory. It is by far the most abundant oxysterol in the brain after parturition, but during development, many more many oxysterols are produced. 24S‑hydroxycholesterol is responsible for 98-99% of the turnover of cholesterol in the central nervous system, which is the source of most of this oxylipin found in plasma. A small amount of it is converted in the brain directly into to 7α,24S‑dihydroxycholesterol by the cytochrome CYP39A1 and thence via side-chain oxidation in peroxisomes to bile acids, such as cholestanoic acid. It is evident that the blood-brain barrier is crossed by constant passive fluxes of oxysterols, but not of cholesterol per se, as a result of their permissive chemical structures and following their concentration gradients. In plasma, it is transported via high-density lipoproteins, as discussed further below. In contrast to humans, CYP46A1 is present in the liver of rodents as well as the brain. 25-Hydroxycholesterol is a relatively minor but biologically important cholesterol metabolite, which is produced rapidly by immune cells during the inflammation resulting from bacterial or viral infections. The dioxygenase enzyme cholesterol 25‑hydroxylase (CH25H in humans), which utilizes a diiron cofactor to catalyze hydroxylation, is the most important route to this metabolite in vivo, although at least two cytochrome P450 enzymes, CYP27A1 and CYP3A4, can catalyze this conversion to a limited extent. Further oxidation by CYP7B1 is a second route to 7α,25‑dihydroxycholesterol, and hence to further oxysterols. 24(S),25-Epoxycholesterol is not produced by the pathways described above but is synthesized in a shunt of the mevalonate pathway using the same enzymes that produce cholesterol, specifically squalene mono-oxygenase and lanosterol synthase, by means of which a second epoxy group is introduced on the other end of squalene from the initial epoxidation. A further mechanism in the brain is the action of CYP46A1 on desmosterol, another intermediate in cholesterol biosynthesis. The oxysterols formed by both autoxidation and enzymatic routes can undergo further oxidation-reduction reactions, and they can be modified by many of the enzymes involved in the metabolism of cholesterol and steroidal hormones, such as esterification and sulfation of position 3, as illustrated for 7-keto-cholesterol as an example in Figure \(32\). In most tissues, esterification of the 3β-hydroxyl group only occurs and requires the activity of sterol O-acyltransferases 1/2 (SOAT1/2 or ACAT1/2) with the participation of cytosolic phospholipase A2 (cPLA2α) to liberate the required fatty acids from phospholipids. In plasma, oxysterols can be esterified by the lecithin–cholesterol acyltransferase (LCAT) for transport in lipoproteins, but in this instance, a diester can be produced from 27‑hydroxycholesterol specifically. Whether such esters are an inert storage form for oxysterols to be liberated on demand by esterases remains to be determined. It is noteworthy that the important human pathogen, Mycobacterium tuberculosis, utilizes a cytochrome P450 enzyme (CYP125) to catalyze C26/C27 hydroxylation of cholesterol as an essential early step in its catabolism as part of the infective process. Catabolism: Because of their increased polarity relative to cholesterol, oxysterols produced by both enzymatic and non-enzymatic means can exit cells relatively easily. A proportion is oxidized further and converted to bile acids, and some are converted to sulfate esters (especially at the 3-hydroxyl group) or glucuronides (see below) for elimination via the kidneys. Non-Enzymatic Oxidation of Cholesterol In biological systems in which both cholesterol and fatty acids are present, it would be expected that autoxidation of polyunsaturated fatty acids by free radical mechanisms would be favored thermodynamically with the formation of isoprostanes from arachidonic acid in phospholipids. However, there are circumstances that can favor cholesterol oxidation in vivo, and, for example, the concentration of cholesterol in low-density lipoprotein particles (LDL) is about three times higher than that of phospholipids, and the rate of cholesterol-hydroperoxide formation can be higher than that of phospholipid hydroperoxides. The rate and specificity of the reaction can depend also on whether it is initiated by free radical species, such as those arising from the superoxide/hydrogen peroxide/hydroxyl radical system (Type I autoxidation), or whether it occurs by non-radical but highly reactive oxygen species such as singlet oxygen, HOCl or ozone (Type II autoxidation). As examples of the main types of products of non-enzymatic oxidation, the structures of a few of the more important of these oxysterols are illustrated in Figure \(33\). Oxysterols produced by this means can vary in the type (hydroperoxy, hydroxy, keto, epoxy), number and position of the oxygenated functions introduced, and nature of their stereochemistry. Derivatives with the A and B rings and the iso-octyl side-chain oxidized are illustrated, but compounds with oxygen groups in position 15 (D ring) are also important biologically. Many are similar to those produced by enzymatic means, although the stereochemistry will usually differ. Like the enzymic products, they are named according to their relationship to cholesterol, rather than by the strict systematic terminology. Mechanisms of autoxidation have been studied intensively in terms of unsaturated fatty acids, and it appears that similar mechanisms operate with sterols. The first event in lipid peroxidation by a radical mechanism is an initiation reaction in which a carbon with a labile hydrogen undergoes hydrogen abstraction by reaction with a free radical, which can be a non-lipid species such as a transition metal or hydroxyl or peroxynitrile radical, and this is followed by oxygen capture. The resulting reactive species recruits further non-oxidized lipids and starts a chain reaction termed the propagation phase. Finally, the reaction is terminated by the conversion of hydroperoxy intermediates to more stable hydroxy products by reaction with endogenous antioxidants such as tocopherols. As an example, the reaction mechanism leading to the production of 7-oxygenated cholesterol derivatives is illustrated in Figure \(34\). Figure \(34\): Cholesterol non-enzymatic oxidation mechanisms In aqueous dispersions, oxidation is initiated by a radical attack from a reactive-oxygen species such as a hydroxyl radical with the abstraction of hydrogen from the C-7 position to form a delocalized three-carbon allylic radical, which reacts with oxygen to produce 7α‑hydroperoxycholesterol, which gradually isomerizes to the more thermodynamically stable 7β-hydroperoxycholesterol. Subsequent enzymic and non-enzymic reactions lead to the 7-hydroxy and 7-keto analogs, which tend to be the most abundant non-enzymatically generated oxysterols in tissues, often accompanied by epoxy-ene and ketodienoic secondary products. Reaction with singlet oxygen (1O2) produces 5α‑hydroperoxycholesterol mainly together with some 6α- or 6β-hydroperoxycholesterol. The reaction does not occur readily at the other allylic carbon 4, presumably because of steric hindrance. When cholesterol is in the solid state, the reaction occurs primarily at the tertiary carbon-25, though some products oxygenated at C-20 may also be produced. Cholesterol hydroperoxides can be converted to stable diols only by the phospholipid hydroperoxide glutathione peroxidase - type 4 (GPx4) and then relatively slowly, but not by the type 1 glutathione peroxidase (GPx1) when in a membrane-bound state. However, in mammalian cells, monomeric GPx4 (~20 kDa), although present in several cellular compartments, including mitochondria, is much less abundant than tetrameric GPx1. Phospholipid-hydroperoxides are reduced most rapidly followed by cholesterol 6β-OOH > 7α/β-OOH >> 5α-OOH. The result is that cholesterol hydroperoxides are expected to have a relatively long half-life and so can potentially be rather dangerous in biological systems. Of these, 5α-OOH with the lowest reduction rate is the most cytotoxic of the hydroperoxides, unfortunately. Epimeric 5,6-epoxy-cholesterols may be formed by a non-radical reaction involving the non-enzymatic interaction of a hydroperoxide with the double bond, a process that is believed to occur in macrophages especially and in low-density lipoproteins (LDL). In this instance, the initial peroxidation product is a polyunsaturated fatty acid; the hydroperoxide transfers an oxygen atom to cholesterol to produce the epoxide, and in so doing is reduced to a hydroxyl. Other non-radical oxidation processes include reaction with singlet oxygen, which is especially important in the presence of light and photosensitizers and can generate 5-hydroxy- as well as 6- and 7-hydroxy products. In addition, the reaction with ozone in the lung can generate a family of distinctive oxygenated cholesterol metabolites. Similarly, a diverse range of oxidation products are generated by peroxidation of the cholesterol and vitamin D precursor 7‑dehydrocholesterol, which has the highest propagation rate constant known for any lipid toward free-radical chain oxidation, and these metabolites have important biological properties. Oxysterols occur in tissues both in the free state and esterified with long-chain fatty acids. For example, in human atherosclerotic lesions, 80–95% of all oxysterols are esterified. Appreciable amounts of oxysterols can be present in foods, especially those rich in cholesterol such as meat, eggs, and dairy products, where they are most probably generated non-enzymically during cooking or processing when such factors as temperature, oxygen, light exposure, the associated lipid matrix, and the presence of antioxidants and water all play a part. Those present in foods can be absorbed from the intestines and transported into the circulation in chylomicrons, but the extent to which dietary sources contribute to tissue levels either of total oxysterols or of individual isomers is not known and is probably highly variable but relatively lower than of cholesterol per se. Oxysterols – Biological Activity General Functions: In tissues in vivo, the very low oxysterol:cholesterol ratio means that oxysterols have little impact on the primary role of cholesterol in cell membrane structure and function, although it has been claimed that oxysterols could cause packing defects and thence atheroma formation in vascular endothelial cells. It is often argued that there are few reliable measurements of cellular or subcellular oxysterol concentrations, because of the technical difficulties in the analysis of the very low concentrations of oxysterols in the presence of a vast excess of native cholesterol; the average levels of 26-, 24- and 7α-hydroxy-cholesterol in human plasma that are often quoted are 0.36, 0.16 and 0.14 μM, respectively. Autoxidation products of cholesterol, especially 7-keto- and 7-hydroxy-cholesterol, are cytotoxic and may be useful markers of oxidative stress or for monitoring of the progression of various diseases. However, experts in the field caution that it can be difficult to extrapolate from experiments in vitro to the situation in vivo, because of the rapidity with which cholesterol can autoxidize in experimental systems and because of the difficulty of carrying out experiments with physiological levels of oxysterols. Nonetheless, aside from their role as precursors of bile acids and some steroidal hormones, it is evident that oxysterols have a variety of roles in terms of maintaining cholesterol homeostasis and perhaps in signaling, where those formed enzymatically are most important. They can exert potent biological effects at physiologically relevant concentrations by binding to various receptors to elicit transcriptional programs, i.e., to regulate gene and hence protein expression. Among many cell membrane receptors for oxysterols to have been identified, nuclear receptors are especially important and include the liver X receptors (LXRs), retinoic acid receptor-related orphan receptors (RORs), estrogen receptors (ERs), and glucocorticoid receptors (GRs). In addition, N-methyl-D-aspartate receptors (NMDARs) are expressed in nerve cells and work over a short time scale to regulate excitatory synaptic function, while G protein-coupled receptors operate at cell membranes and are activated by molecules outside the cell to activate signaling pathways within the cell. As various isoforms of these receptors exist in different tissues, and these can interact with several oxysterols, only a brief summary of this complex topic is possible here. A family of oxysterol-binding proteins (OSBP) transports and regulates the metabolism of sterols and targets oxysterols to specific membranes and especially to contact sites between organelles with the transport of phosphatidylinositol 4-phosphate in the reverse direction (see our web page on the latter). In this way, they can enable oxysterols to regulate membrane composition and function and mediate intracellular lipid transport. As with cholesterol, oxysterols can be eliminated from cells by transporters such as the ATP-binding cassette proteins ABCA1 and ABCG1, and they are transported in the blood-stream within lipoproteins, especially in association with HDL and LDL and mainly in the esterified form. Cholesterol homeostasis: While cholesterol plays a key role in its own feedback regulation, there is some evidence that oxysterols are regulators of cholesterol concentration in cell membranes, and that 25‑hydroxycholesterol and 24(S),25‑epoxycholesterol may be especially effective, although the other side-chain oxysterols 22-, 24- and 27‑hydroxycholesterol have been implicated. Several mechanisms appear to be involved, and it is suggested that 24(S),25‑epoxycholesterol especially acts as a ligand for the liver X receptor, which forms a heterodimer with the retinoic X receptor, to inhibit the transcription of key genes in cholesterol biosynthesis, as well as directly inhibiting or accelerating the degradation of such important enzymes in the process as HMG-CoA reductase and squalene synthase. Similarly, both 26-hydroxylanosterol and 25-hydroxycholesterol inhibit HMG-CoA reductase. 25‑Hydroxycholesterol inhibits the transfer of the 'sterol regulatory element binding protein' (SREBP-2) to the Golgi for processing to its active form as a transcription factor for the genes of the cholesterol biosynthesis pathway, and it stimulates the enzyme acyl-CoA:cholesterol acyl transferase in the endoplasmic reticulum to esterify cholesterol. By such mechanisms, these oxysterols fine-tune cholesterol homeostasis and ensure smooth regulation rather than substantial fluctuations in tissue concentration. Oxysterols and the immune system: Oxysterols and especially are known to have vital and diverse roles in immunity by regulating both the adaptive and innate immune responses to infection. For example, infection with viruses leads to the production of type I interferon, and in macrophages, this induces synthesis of 25‑hydroxycholesterol, which in general is regarded as anti-inflammatory and exerts broad antiviral activity by activating integrated stress response genes and reprogramming protein translation again via its interaction with LXR receptors. It is a potent inhibitor of SARS-CoV-2 replication, for example, possibly by a mechanism involving the blocking of cholesterol export from the late endosome/lysosome compartment and depletion of membrane cholesterol levels. However, the formation of 25‑hydroxycholesterol may be harmful in the case of influenza infections, as it can lead to over production of inflammatory metabolites. Similarly, the biosynthesis of 25-hydroxycholesterol in macrophages is stimulated by the endotoxin Kdo2-lipid A, the active component of the lipopolysaccharide present on the outer membrane of Gram-negative bacteria, which acts as an agonist for Toll-like receptor 4 (TLR4). There is enhanced expression of the oxygenase CH25H in immune cells in response to bacterial and viral infection. Many oxysterol species are active in a range of immune cells subsets, mediated through the control of LXR and SREBP signaling, but also by acting as ligands for RORs, and for the cell surface receptors G protein-coupled receptor 183 (GPR183) or CXCR2. Activation of LXR tends to dampen the immune response. In response to various stimuli, they can operate through ion channels to effect rapid changes in intracellular ion concentrations, especially of Ca2+, to bring about changes in membrane potential, cell volume, cell death (apoptosis, autophagy, and necrosis), gene expression, secretion, endocytosis, or motility. For example, 27‑hydroxycholesterol in human milk is reported to be active against the pathogenic human rotavirus and rhinovirus of importance in pediatrics, and 7-Dehydrocholesterol has anti-viral properties also. While they can exert their immune functions within the cell in which they are generated, oxysterols can also operate in a paracrine fashion towards other immune cells. 25‑Hydroxycholesterol in particular can have either pro- or anti-inflammatory effects, depending upon the conditions, but the enzyme CH25H responsible for its biosynthesis is induced markedly in macrophages activated by inflammatory agents. It is reported to have a regulatory effect on the biosynthesis of sphingomyelin, which is required with cholesterol for the formation of raft sub-domains in membranes, where signaling molecules are concentrated, and together with other oxysterols, such as 24S,25-epoxycholesterol, to regulate the activities of the hedgehog proteins involved in embryonic development. Metabolites of 25‑hydroxycholesterol, such as 7α,25‑dihydroxycholesterol, and further oxidation products, are pro-inflammatory act as chemoattractants to lymphocytes; they have a role in the regulation of immunity in secondary lymphoid organs by interactions with the receptor GPR183. Oxysterols in brain: Oxysterols are especially important for cholesterol homeostasis in the brain, which contains 25% of the total body cholesterol, virtually all of it in unesterified form, in only about 2% of the body volume. Cholesterol is a major component of the plasma membrane especially, where it serves to control fluidity and permeability. This membrane is produced in large amounts in the brain and is the basis of the compacted myelin, which is essential for the conductance of electrical stimuli and contains about 70% of brain cholesterol. While this pool is relatively stable, the remaining 30% is present in the membranes of neurons and glial cells of gray matter and is more active metabolically. Even in the fetus and the newborn infant, all the cholesterol required for growth is produced by synthesis de novo in the brain, not by transfer from the circulation across the blood-brain barrier, which consists of tightly opposed endothelial cells lining the extensive vasculature of the tissue. The fact that this pool of cholesterol in the brain is independent of circulating levels must reflect a requirement for constancy in the content of this lipid in membranes and myelin. In adults, although there is a continuous turnover of the membrane, the cholesterol is efficiently re-cycled and has a remarkably high half-life (up to 5 years). The rate of cholesterol synthesis is a little greater than the actual requirement so net movement of cholesterol out of the central nervous system must occur. An important component of this system is apolipoprotein E (Apo E), a 39-kDa protein, which is highly expressed in the brain and functions in the cellular transport of cholesterol and in cholesterol homeostasis. Apo E complexes with cholesterol are required for transport from the site of synthesis in astrocytes to neurons. Hydroxylation by CYP46A1 of cholesterol to 24(S)‑hydroxycholesterol (cerebrosterol) is responsible for 50–60% of all cholesterol metabolism in the adult brain. If cholesterol itself cannot cross the blood-brain barrier, this metabolite is able to do so with relative ease. When the hydroxyl group is introduced into the side chain, this oxysterol causes a local re-ordering of membrane phospholipids such that it is more favorable energetically to expel it at a rate that is orders of magnitude greater than that of cholesterol per se, though still only 3-7 mg per day. There is a continuous flow of the metabolite from the brain into the circulation, much of it in the form of the inactive sterol ester, where it is transported by lipoprotein particles to the liver for further catabolism, i.e., it is hydroxylated in position 7 and then converted to bile acids. This is illustrated in Figure \(35\). Both 24(S)-hydroxycholesterol and 24(S),25-epoxycholesterol are believed to be important in regulating cholesterol homeostasis in the brain. They interact with the specific nuclear receptors involved in the expression and synthesis of proteins involved in sterol transport, and for example, 24‑hydroxy-cholesterol regulates the transcription of Apo E. In particular, it is an agonist of the nuclear liver X receptors (LXRs), influencing the expression of those LXR target genes involved in cholesterol homeostasis and inflammatory responses. It is also a high-affinity ligand for the retinoic acid receptor-related orphan receptors α and γ (RORα and RORγ). In this way, it can act locally to affect the functioning of neurons, astrocytes, oligodendrocytes, and vascular cells. 24(S)-Hydroxycholesterol down-regulates the trafficking of the amyloid precursor protein and may be a factor in preventing neurodegenerative diseases. Especially high levels of 24(S)‑hydroxycholesterol are observed in the plasma of human infants and in patients with brain trauma, while reduced levels are found in the plasma of patients with neurodegenerative diseases, including Parkinson’s disease, multiple sclerosis, and Alzheimer's disease. In contrast, there are elevated levels in the brain and especially cerebrospinal fluid in patients with these conditions, where it may be a marker of neurodegeneration. Increased expression of cholesterol 24-hydroxylase (CYP46A1) is believed to improve cognition, while a reduction leads to poor cognitive performance, as occurs at advanced stages of the disease, probably reflecting a selective loss of neuronal cells, and it may be a factor in age-related macular degeneration. An excess of 24(S)‑hydroxycholesterol and especially of its ester form can lead to neuronal cell death, and elevated levels in plasma are reported to be a potential marker for Autism Spectrum Disorders in children. On the other hand, it may be protective against glioblastoma, the most common primary malignant brain tumour in adults via activation of LXRs. 27‑Hydroxy-cholesterol diffuses across the blood-brain barrier from the bloodstream into the brain (in the reverse direction to 24‑hydroxycholesterol), where it does not accumulate but is further oxidized and then exported as steroidal acids. This flux may regulate certain key enzymes within the brain, and there are suggestions that the balance between the levels of 24- and 27-hydroxy-cholesterol, especially excess of the latter, may be relevant to the generation of β-amyloid peptides in Alzheimer's disease by reducing insulin-mediated glucose uptake by neurons. While 7β-hydroxycholesterol is pro-apoptotic, any links with Alzheimer's disease are unproven although there is a school of thought that other oxidized cholesterol metabolites may be major factors behind the development of this disease. For example, seco-sterols such as 3β‑hydroxy-5-oxo-5,6-secocholestan-6-al and its stable aldolization product, the main ozonolysis metabolites derived from cholesterol, have been detected in brain samples of patients who have died from Alzheimer's disease and Lewy body dementia; they are also found in atherosclerotic lesions. Oxidation products of the cholesterol precursor 7‑dehydrocholesterol and especially 3β,5‑dihydroxycholest-7-ene-6-one are involved in the pathophysiology of the human disease Smith-Lemli-Opitz syndrome. Cell differentiation: Oxysterols can influence the differentiation of many cell types and this was first studied in the skin, where 22(R)- and 25(R)‑hydroxycholesterol were shown to induce human keratinocyte differentiation. Subsequently, by stimulating nuclear binding receptors, oxysterols were found to have similar effects on mesenchymal stem cells. There have been many reports of the involvement of oxysterols in disease processes, especially atherosclerosis and the formation of human atherosclerotic plaques, but also cytotoxicity, necrosis, inflammation, immuno-suppression, phospholipidosis and gallstone formation. They have been implicated in the development of cancers, especially those of the breast, prostate, colon, and bile duct. For example, 27‑hydroxycholesterol is an element in cholesterol elimination from macrophages and arterial endothelial cells, but it is also an endogenous ligand for the human nuclear estrogen receptor (ERα) and the liver X receptor, and it modulates their activities with effects upon various human disease states, including cardiovascular dysfunction and progression of cancer of the breast and prostate, as well as having an involvement in the regulation of bone mineralization (osteoporosis). It has been linked to cancer metastasis through effects on immune cells, and there is hope that pharmacological inhibition of CYP27A1 and thence the formation of 27‑hydroxycholesterol may be a useful strategy in the treatment of breast cancer; CYP7A1 gene polymorphism has been associated with colorectal cancer. In contrast, oxysterols can interfere in the proliferation of several types of cancer cell (glioblastoma, leukemia, colon, breast, and prostate cancer). Cholesterol 5,6-epoxide (with either 5α or 5β stereochemistry) is formed non-enzymatically in tissues, but it is also believed to be produced by an unidentified cytochrome P450 enzyme in the adrenal glands. While it was for some time believed to be a causative agent in cancer, it is now recognized that downstream metabolites are responsible. Thus, cholesterol epoxide hydrolase converts cholesterol 5,6-epoxide into cholestane-3β,5α,6β-triol, which is transformed by 11β‑hydroxysteroid-dehydrogenase-type-2 into the oncometabolite 3β,5α-dihydroxycholestan-6-one (oncosterone). By binding to the glucocorticoid receptor, this oncosterone stimulates the growth of breast cancer cells, and it also acts as a ligand to the LXR receptors, which may mediate its pro-invasive effects. In contrast, in normal breast tissue, cholesterol 5,6‑epoxide is metabolized to the tumor suppressor metabolite, a steroidal alkaloid designated dendrogenin A that is a conjugation product with histamine and controls a nuclear receptor to trigger lethal autophagy in cancers; its synthesis is greatly reduced in cancer cells. Tamoxifen, a drug that is widely used against breast cancer, binds to the cholesterol 5,6-epoxide hydrolase, which is also a microsomal anti-estrogen binding site (AEBS), to inhibit its activity. 7-Ketocholesterol is a major oxysterol produced during the oxidation of low-density lipoproteins, and is one of the most abundant in plasma and atherosclerotic lesions; it accumulates in erythrocytes of heart failure patients. It has a high pro-apoptotic potential and associates preferentially with membrane lipid raft domains. As it is not readily exported from macrophages, it impairs cholesterol efflux and promotes the foam cell phenotype. In cardiomyocytes, this accumulation can lead to cell hypertrophy and death, and it has been suggested that oxysterols are a major factor precipitating morbidity in atherosclerosis-induced cardiac diseases and inflammation-induced heart complications. Photoxidation in the retina via the action of free radicals or singlet oxygen generates unstable cholesterol hydroperoxides, which may be involved in age-related macular degeneration. For example, these compounds can quickly be converted to highly toxic 7α- and 7β‑hydroxycholesterols and 7‑ketocholesterol, depending on the status of tissue oxidases and reductases. Three separate enzymatic pathways have developed in the eye to neutralize their activities. These sterols are metabolized by novel branches of the acidic pathway of bile acid biosynthesis. Those oxysterols formed non-enzymatically can be most troublesome in relation to disease in general. For example, they are enriched in pathologic cells and tissues, such as macrophage foam cells, atherosclerotic lesions, and cataracts. They may regulate some of the metabolic effects of cholesterol, but as cautioned above, effects observed in vitro may not necessarily be of physiological importance in vivo. Various oxysterols have been implicated in the differentiation of mesenchymal stem cells and the signaling pathways involved in this process. High levels of 7‑hydroxycholesterol and cholestane-3β,5α,6β-triol are characteristic of the lysosomal storage diseases Niemann-Pick types B and C and of lysosomal acid lipase deficiency. Cholesterol hydroperoxides: With the aid of START domain proteins, cholesterol hydroperoxides can translocate from a membrane of origin to another membrane such as mitochondria. Such transfer of free radical-generated 7-hydroperoxycholesterol, for example, has adverse consequences in that there is impairment of cholesterol utilization in steroidogenic cells, and of anti-atherogenic reverse-cholesterol transport in vascular macrophages. The antioxidant activity of GPx4 may be crucial for the maintenance of mitochondrial integrity and functionality in these cells. Vitamin D Vitamin D encompasses two main sterol metabolites that are essential for the regulation of calcium and phosphorus levels and thence for bone formation in animals. However, these have many other functions, including induction of cell differentiation, inhibition of cell growth, immunomodulation, and control of other hormonal systems. Vitamin D (with calcium) deficiency is responsible for the disease rickets in children in which bones are weak and deformed, and it is associated with various cancers and autoimmune diseases. Ultraviolet light mediates cleavage of 7-dehydrocholesterol, an important intermediate in the biosynthesis of cholesterol, with the opening of the second (B) ring in the skin to produce pre-vitamin D, which rearranges spontaneously to form the secosteroid vitamin D3 or cholecalciferol. Its structure is shown in Figure \(36\). The newly generated vitamin D3 is transported to the liver where it is subject to 25-hydroxylation and thence to the kidney for 1α-hydroxylation to produce the active form 1α,25-dihydroxyvitamin D3 (calcitriol); this is a true hormone and serves as a high-affinity ligand for the vitamin D receptor in distant tissues. For transportation in plasma, it is bound to a specific glycoprotein termed unsurprisingly, the 'vitamin D binding protein (BDP)'. Vitamin D2 or ergocalciferol is derived from ergosterol, which is obtained from plant and fungal sources in the diet. Vitamin D3 functions by activating a cellular receptor (vitamin D receptor or VDR), a transcription factor binding to sites in the DNA called vitamin D response elements. There are thousands of such binding sites, which together with co-modulators regulate innumerable genes in a cell-specific fashion. In this way, it enhances bone mineralization by promoting dietary calcium and phosphate absorption, as well as having direct effects on bone cells. In addition, it functions as a general development hormone in many different tissues, while together with Vitamin D2 it has profound effects on immune responses in the defense against microbes. Steroidal Hormones and their Esters Steroidal hormones cannot be discussed in depth here as their structures, biosynthesis, and functions comprise a rather substantial and specialized topic. In brief, animal tissues produce small amounts of vital steroidal hormones from cholesterol as the primary precursor with 22R-hydroxycholesterol, produced by hydroxylation by the cholesterol side-chain cleavage enzyme (P450scc), as the first of its metabolites in the pathway. This step involves the 'STAR' protein which enables the transport of cholesterol into mitochondria where conversion to pregnenolone is rate-limiting and involves first hydroxylation and then cleavage of the side-chain. After export from the mitochondria, this can be converted directly to progesterone or in several steps to testosterone. 17β-Estradiol, for example, is the most potent and important of the endogenous estrogens; it is made mainly in the follicles of the ovaries and regulates menstrual cycles and reproduction, but it is also present in testicles, adrenal glands, fat, liver, breasts, and brain. Testosterone is the primary male sex hormone and an anabolic steroid, and it is produced mainly in the testes; it has a key function in the development of male reproductive tissues such as testes and prostate, in addition to promoting secondary sexual characteristics. Pregnane neurosteroids are synthesized in the central nervous system. Cholesterol homeostasis is therefore vital to fertility and a host of bodily functions. The structures of key steroidal hormones are shown in Figure \(37\). Steroidal esters accumulate in tissues such as the adrenal glands, which synthesize corticosteroids such as cortisol and aldosterone and are responsible for releasing hormones in response to stress and other factors. It is also apparent that fatty acyl esters of estradiols, such as dehydroepiandrosterone, accumulate in adipose tissue in post-menopausal women. Small amounts of estrogens acylated with fatty acids at the C-17 hydroxyl are present in the plasma lipoproteins. In each instance, they appear to be biologically inert storage or transport forms of the steroid. Eventually, esterified steroids in low-density lipoproteins (LDL) particles are taken up by cells via lipoprotein receptors, and then are hydrolyzed to release the active steroid. Pharmaceutical interest in oleoyl-estrone, a naturally occurring hormone in humans, which was found to induce a marked loss of body fat while preserving protein stores in laboratory animals, has declined as clinical trials with humans were not successful. Sterols 3. Sterols and their Conjugates from Plants and Lower Organisms Plant Sterols - Structures and Occurrence Plants, algae, and fungi contain a rather different range of sterols from those in animals. Like cholesterol, to which they are related structurally and biosynthetically, plant sterols form a group of triterpenes with a tetracyclic cyclopenta[a]phenanthrene structure and a side chain at carbon 17, sometimes termed the C30H50O triterpenome. The four rings (A, B, C, D) have trans ring junctions, and the side chain and two methyl groups (C-18 and C-19) are at an angle to the rings above the plane with β stereochemistry (as for a hydroxyl group commonly located on C-3 also). The basic sterol from which other sterol structures are defined is 5α-cholestan-3β-ol with the numbering scheme recommended by IUPAC as shown in Figure \(38\). The phytosterols (as opposed to zoosterols) include campesterol, β-sitosterol, stigmasterol, and Δ5‑avenasterol, some of which are illustrated in Figure \(39\). These more common plant sterols have a double bond in position 5, and a definitive feature – a one- or two-carbon substituent with variable stereochemistry in the side chain at C-24, which is preserved during subsequent metabolism. For example, campesterol is a 24-methylsterol, while β-sitosterol and stigmasterol are 24‑ethylsterols. Occasionally, there is a double bond in this chain that can be of the cis or trans configuration as in stigmasterol (at C22) or fucosterol (C24), the main sterol in green algae. Phytosterols can be further classified on a structural or biosynthetic basis as 4‑desmethyl sterols (i.e. with no substituent on carbon‑4), 4α‑monomethyl sterols and 4,4‑dimethyl sterols. The most abundant group is the 4‑desmethyl sterols, which may be subdivided into Δ5-sterols (illustrated above), Δ7‑sterols (e.g. α-spinasterol) and Δ5,7-sterols depending on the position of the double bonds in the B ring. As the name suggests, brassicasterols (24‑methyl-cholesta-5,22-dien-3β-ol and related sterols) are best known from the brassica family of plants, but they are also common constituents of marine algae (phytoplankton). Phytostanols (fully saturated) are normally present at trace levels only in plants, but they are relatively abundant in cereal grains. Many different sterols may be present in photosynthetic organisms, and the amounts and relative proportions are dependent on the species. Over 250 different phytosterols have been recorded with 60 in corn (maize) alone, for example. As a rough generality, a typical plant sterol mixture would be 70% sitosterol, 20% stigmasterol, and 5% campesterol (or >70% 24-ethyl-sterols and <30% 24-methyl-sterols), although this will vary with the stage of development and in response to stress. Table 1 contains data on the main components from some representative commercial seed oils. Table \(2\). Sterol composition in some seed oils of commercial importance (mg/Kg). corn cottonseed olive palm rapeseed safflower soybean sunflower cholesterol - - - 26 - - - - campesterol 2691 170 28 358 1530 452 720 313 stigmasterol 702 42 14 204 - 313 720 313 β‑sitosterol 7722 3961 1310 1894 3549 1809 1908 2352 Δ5‑avenasterol 468 85 29 51 122 35 108 156 Δ7‑stigmasterol 117 - 58 25 306 696 108 588 Δ7‑avenasterol - - - - - 104 36 156 brassicasterol - - - - 612 - - - other - - - - - 69 - 39 Data adapted from Gunstone, F.D. et al. The Lipid Handbook (Second Edition) (Chapman & Hall, London) (1994). Cholesterol is usually a minor component only of plant sterols (<1%), but it is unwise to generalize too much as it can be the main sterol component of red algae and of some families of higher plants such as in the Solanaceae, Liliaceae and Scrophylariaceae. It can also be a significant constituent sterol of chloroplasts, shoots, pollen and leaf surface lipids in other plant families; wheat roots contain 10% and Arabidopsis cells 19% of the sterols as cholesterol. Yeasts and fungi tend to contain ergosterol as their main sterol (see below). Ecdysteroids (phytoecdysteroids) are polyhydroxylated plant sterols that can occur in appreciable amounts in some species. Sterols are also found in some bacterial groups but not in archaea, and hopanoids in bacteria are considered to be functional triterpenic counterparts. Sterols can occur in plants in the 'free' state, i.e. in which the sterol hydroxyl group is not linked to any other moiety, but they are usually present also as conjugates with the hydroxyl group covalently bound via an ester bond to a fatty acid, for example, i.e. as sterol esters, or via a glycosidic linkage to glucose (and occasionally other sugars), i.e. as steryl glycosides. Plant Sterols - Biosynthesis The biosynthetic route to plant sterols resembles that to cholesterol in many aspects in that it follows an isoprenoid biosynthetic pathway with isopentenyl pyrophosphate, derived primarily from mevalonate, as the key building block in the cytoplasm (but not plastids) at least. The main pathway for the biosynthesis of isopentenyl pyrophosphate and dimethylallyl pyrophosphate, the isoprene units, is described previously and so need not be repeated here. It is known as the 'mevalonic acid (MVA) pathway' and functions in the cytosol, endoplasmic reticulum and mitochondria. However, an alternative pathway that does not use mevalonic acid as a precursor was established first for bacterial hopanoids, but has since been found in plant chloroplasts, algae, cyanobacteria, eubacteria, and some parasites (but not in animals). This route is variously termed the ‘non-mevalonate’, ‘1‑deoxy-D-xylulose-5-phosphate’ (DOXP) or better the 2C-methyl-D-erythritol 4-phosphate (MEP) pathway as the last compound is presumed to be the first committed intermediate in sterol biosynthesis by this route. In the first step, pyruvate and glyceraldehyde phosphate are combined to form deoxyxylose phosphate, which is in turn converted to 2C-methyl-D-erythritol 4-phosphate. The pathway then proceeds via various erythritol intermediates until isopentenyl pyrophosphate and dimethylallyl pyrophosphate are formed. This pathway is illustrated in Figure \(40\). There is evidence that some of the isoprene units are exchanged between the cytoplasm and plastids. In much of the plant kingdom, both the MVA and MEP pathways operate in parallel, but green algae use the MEP pathway only. Thereafter, sterol biosynthesis continues via squalene and (3S)-2,3-oxidosqualene. In photosynthetic organisms (as opposed to yeast and fungi), the subsequent steps in the biosynthesis of plant sterols differ from that for cholesterol in that the important intermediate in the route from squalene via 2,3-oxidosqualene is cycloartenol, rather than lanosterol, and this is produced by the action of a 2,3(S)‑oxidosqualene-cycloartenol cyclase (cycloartenol synthase). Then, the enzyme sterol methyltransferase 1 is of special importance in that it converts cycloartenol to 24-methylene cycloartenol, as the first step in introducing the methyl group onto C-24, while the enzyme cyclopropyl sterol isomerase is required to open the cyclopropane ring. Animals lack the sterol C24-methyltransferase gene. While this pathway is in essence linear up to the synthesis of 24-methylene lophenol, a bifurcation then occurs that results in two alternative pathways, one of which leads to the synthesis of sitosterol and stigmasterol and the other to that of campesterol. This pathway is shown in Figure \(41\). In fact, there are more than thirty enzyme-catalyzed steps in the overall process of plant sterol biosynthesis, each associated with membranes, and detailed descriptions are available from the reading list below. The 4,4-dimethyl- and 4α-methylsterols are part of the biosynthetic pathway, but are only minor if ubiquitous sterol components of plants. New biosynthetic pathways are now being discovered by genome analysis that reveal the complexity of sterol biosynthesis in different plant species. Dinoflagellates produce a characteristic 4-methylsterol termed dinosterol and others like gorgosterol via lanosterol as precursor. Protozoans synthesize many different sterols related to those in plants. For example, some species of Acanthamoeba and Naegleria produce both lanosterol and cycloartenol, but only the latter is used for the synthesis of other sterols, especially ergosterol, but in other protozoan species, sterol biosynthesis occurs via lanosterol. The best studied bacterial pathway is that of the methylotroph Methylococcus capsulatus, which produces a number of unique Δ18(14)-sterols and is known to possess a squalene epoxidase and a lanosterol-14-demethylase. Cholesterol in plants is produced from cycloartenol as the key intermediate with the Sterol Side Chain Reductase 2 (SSR2) as the key enzyme. It is now established that the cholesterol biosynthetic pathway in tomato plants comprises 12 enzymes acting in 10 steps. Of these, half evolved through gene duplication and divergence from phytosterol biosynthetic enzymes, whereas others act reciprocally in both cholesterol and phytosterol metabolism. Algae can also produce cholesterol in a multi-step process from cycloartenol, and many more sterols via 24-methylene lophenol as the key intermediate. It is hoped that genetic manipulation of these enzymes will lead to plants that synthesize high-value steroidal products. Oxidation: Phytosterols can be subjected to non-enzymatic oxidation with the formation of oxysterols in a similar manner to that of cholesterol in animals, resulting in ring products such as hydroxy-, keto-, epoxy- and triol-derivatives, and further enzymic reactions can oxidize the side chain. However, photosensitized oxidation is more common in plants and is much faster (>1500 times); it starts with the ene-addition of singlet oxygen (1O2) on either side of the double bond in the B ring to generate 5α-/6α-/6β-hydroperoxysterols, of which 5α-OOH is the most abundant and rearranges to form the more stable 7α‑OOH isomer. This is the main reaction in foods stored under LED lighting in food retailers. Plant Sterols - Function Like cholesterol, plant sterols are amphiphilic and are vital constituents of all membranes, and especially of the plasma membrane, the mitochondrial outer membrane and the endoplasmic reticulum. The three-dimensional structure of the plant sterols is such that there are planar surfaces at both the top and the bottom of the molecules, which permit multiple hydrophobic interactions between the rigid sterol and the other components of membranes. Indeed, they must determine the physical properties of membranes to an appreciable extent. It is believed that campesterol, β-sitosterol, and 24-methylcholesterol (in this order) are able to regulate membrane fluidity and permeability in plant membranes by restricting the mobility of fatty acyl chains in a similar manner to cholesterol in mammalian cells, but stigmasterol has much less effect on lipid ordering and no effect on the permeability of membranes. In the plasma membrane, plant sterols associate with the glycosphingolipids such as glucosylceramide, and glycosylinositolphosphoceramides in raft-like sub-domains, analogous to those in animal cells, and these support the membrane location and activities of many proteins with important functions in plant cells. The sterol glycosides are especially important in this context (see below). Sterols (and their conjugates) are involved in plant membrane adaptations to changes in temperature and other biotic and abiotic stresses. For example, β‑sitosterol is a precursor of stigmasterol via the action of a C22-sterol desaturase, and the ratio of these two sterols is important to the resistance of A. thaliana plants to low and high temperatures. In addition, plant sterols can modulate the activity of membrane-bound enzymes. Thus, stigmasterol and cholesterol regulate the activity of the Na+/K+-ATPase in plant cells, probably in a manner analogous to that of cholesterol in animal cells. Stigmasterol may be required specifically for cell differentiation and proliferation. As well as being the precursor of plant steroidal hormones, campesterol, is a signaling molecule that regulates growth, development, and stress adaptation. Perhaps surprisingly, cholesterol is a precursor for the biosynthesis of some steroidal saponins and alkaloids in plants, for example, the well-known steroidal glycoalkaloid in potato (α-solanine), as well as of other steroids including the phytoecdysteroids (in some species they are derived from lathosterol). While the physiological roles of ecdysteroids in plants yet to be been confirmed, they are believed to enhance stress resistance by promoting health and vitality. Withanolides are complex oxysterols, which are believed to be defense compounds against insect herbivores. Steroidal Plant Hormones They have crucial importance for plant growth processes, including cell elongation, division, differentiation, immunity, and development of reproductive organs, and they are involved in the regulation of innumerable aspects of metabolism. Via signal transduction pathways, they interact with transcription factors through phosphorylation cascades to regulate the expression of target genes. Brassinosteroids are also signaling molecules in abiotic stress responses such as drought, salinity, high temperature, low temperature, and heavy metal stresses. Outwith plants, they may have biomedical applications as anticancer drugs for endocrine-responsive cancers to induce apoptosis and inhibit growth. Some plant species produce small amounts of steroid hormones that are often considered to be of animal origin only, including progesterone and testosterone, and these may have physiological roles in plants. Sterol Esters in Higher Plants Sterol esters are present in all plant tissues, but they are most abundant in tapetal cells of anthers, pollen grains, seeds, and senescent leaves. In general, they are minor components relative to the free sterols other than in waxes. Usually, the sterol components of sterol esters are similar to the free sterols, although there may be relatively less of stigmasterol. The fatty acid components tend to resemble those of the other plant tissue lipids, but there can be significant differences on occasion. Sterol esters are presumed to serve as inert storage forms of sterols, as they are often enriched in the intermediates of sterol biosynthesis and can accumulate in lipid droplets within the cells. However, they have been found in some membranes, especially in microsomes and mitochondrial preparations, although their function there is uncertain. They may also have a role in transport within cells and between tissues, as they can be present in the form of soluble lipoprotein complexes. Biosynthesis of sterol esters in A. thaliana is known to occur in the endoplasmic reticulum by the action of a phospholipid:sterol acyltransferase, which catalyzes the transfer of a fatty acyl group to the sterol from position sn-2 of phospholipids - mainly phosphatidylethanolamine; the enzyme is very different from those in animals and yeasts. However, an acyl CoA:sterol acyltransferase closer in structure to the animal enzyme has been characterized also; it prefers saturated fatty acyl-CoAs as acyl donors and cycloartenol as the acceptor molecule. The enzymes responsible for the hydrolysis of sterol esters in A. thaliana are not yet known. Certain distinctive phytosterol esters occur in the aleurone cells of cereal grains, including trans-hydroxycinnamate, ferulate (4-hydroxy-3-methoxycinnamate), and p-coumarate esters. Similarly, rice bran oil is a rich source of esters of ferulic acid and a mixture of sterols and triterpenols, termed 'γ-orizanol'’, and an example of one of these compounds is illustrated in Figure \(43\). This is sold as a health food supplement, because of the claimed beneficial effects, including cholesterol-lowering and antioxidant activities, while enhancing muscle growth and sports performance. However, none of these effects have been confirmed by rigorous clinical testing. Sterol Glycosides Leaf and other tissue in plants contain a range of sterol glycosides and sterol acyl-glycosides in which the hydroxyl group at C3 on the sterol is linked to the sugar by a glycosidic bond. Other than in the genus Solanum, where they can represent up to 85% of sterol fraction in tomato fruit as an example, they tend to be minor components relative to other lipids. Typical examples (glucosides of β-sitosterol) are illustrated in Figure \(44\). Most of the common plant sterols occur in this form, but Δ5 sterols are preferred (Δ7 in some genera). Glucose is the most common carbohydrate moiety but galactose, mannose, xylose, arabinose can also be present depending on plant species; occasionally, complex carbohydrates with up to five hexose units linked in a linear fashion are present. Algae also contains sterol glycosides with a wide range of sterol and carbohydrate components. Plant, animal, fungal, and most bacterial steryl glycosides have a β‑glycosidic linkage, but in a few bacterial species there is an α-linkage. Similarly, the nature of the fatty acid component in the acyl sterol glycosides can vary as well as the hydroxyl group to which they are linked, although it is usually position 6 of the glucose moiety. In potato tubers, for example, the 6'-palmitoyl-β-D-glucoside of β-sitosterol is the major species, while the corresponding linoleate derivative predominates in soybeans. Usually, the sterol acyl-glycosides are present at concentrations that are two- to tenfold greater than those of the non-acylated forms. They are known to be located in the plasma membrane, tonoplasts and endoplasmic reticulum. Biosynthesis involves the reaction of free sterols with a glucose unit catalyzed by a sterol glycosyltransferase, or by the reaction of the sterol with uridine diphosphoglucose (UDP-glucose) and UDP-glucose:sterol glucosyltransferase on the cytosolic side of the plasma membrane. The acyl donor for acyl sterol glycoside synthesis is not acyl-coenzyme A but is believed to be a glycerolipid. Steryl β-D-glycoside hydrolases have been characterized from plants that reverse this reaction, but no fatty acyl hydrolase activity for sterol acyl-glycosides is yet known. One route to the biosynthesis of glucosylceramides in plants involves the transfer of the glucose moiety of sterol glycosides to ceramide. The functions of sterol glycosides and sterol acyl-glycosides are slowly being revealed, and they are believed to be significant components of the plasma membrane that associate with sphingolipids in raft-like domains; the esterified form especially may be involved in the adaptation of plant membranes to low temperatures and other stresses. It is possible that they have a role in signal transmission through membranes, and they are reported to be beneficial in the response to pathogens. It seems probable that sterol glycosides are oriented with the sterol moiety buried in the hydrophobic core of the lipid bilayer with the sugar located in the plane of the polar head groups of the membrane, while with sterol acyl-glycosides both the sterol moiety and the fatty acid chain are embedded in the hydrophobic core of the membrane. Sitosterol-β-D-glucoside in the plasma membrane is believed to be the primer molecule for cellulose synthesis in plants, as in cotton (Gossypium arboreum) fiber, where it may be required for the initiation of glucan polymerization. The sterol is eventually removed from the polymer by a specific cellulase enzyme (the multimeric cellulase synthase is believed to be stabilized by sterols in the plasma membrane). Sterol glycosides appear to be essential for the pathogenicity of certain fungi and for some bacteria, and ergosterol glycosides especially are especially troublesome components of plant fungal pathogens. Sterol glycosides have only rarely been reported from organisms other than plants and fungi, although some bacteria, such as the gram-negative bacterium Helicobacter pylori and Borrelia burgdorferi, the causative agent of Lyme disease produce cholesterol glucoside from host cholesterol. On the other hand, cholesteryl glucoside has been found as a natural component of a few animal tissues, and through acting as immunoadjuvants, sterol glycosides are reported to be efficacious in protecting animal hosts against lethal Cryptococcal infections. In the human diet, sterol glycosides have potential benefits in that like free sterols they inhibit the absorption of cholesterol from the gut and reduce the plasma cholesterol levels. The fatty acids are removed from sterol acyl-glycosides by enzymes in the intestine. A number of species of monocotyledons contain complex steroidal saponins, which consist of an aglycone based on a triterpenoid furostanol or spirostanol skeleton (derived from cholesterol) and an oligosaccharide chain of two to five hexose or pentose moieties attached to the 3β-hydroxyl group of the sterol. These can interact with cholesterol in plant membranes to form insoluble complexes, which increase membrane permeability. Ergosterol and Other Sterols in Yeasts and Fungi Yeasts and fungi, together with microalgae and protozoa, can contain a wide range of different sterols. However, ergosterol ((22E)‑ergosta-5,7,22-trien-3β-ol) is the most common sterol in fungi and yeast, and is accompanied by other sterols not normally abundant in higher plants including cholesterol, 24-methyl cholesterol, 24-ethyl cholesterol, and brassicasterol, depending upon species. In Saccharomyces cerevisiae, which is widely studied as a model species of yeasts, ergosterol is the most abundant sterol (ca. 12% of all lipids), with the highest levels in the plasma membrane (up to 40% of the lipids or 90% of the total cell sterols). Its structure is shown in Figure \(45\). Like cholesterol and in contrast to the plant sterols, it is synthesized in the endoplasmic reticulum via lanosterol as the key intermediate and then zymosterol, but the pathway diverges at this stage to produce fecosterol on the way to ergosterol (see the reading list below for further details). Ergosterol is transported to other organelles within the cell in a non-vesicular manner by two families of evolutionarily conserved sterol-binding proteins - 'Osh' and 'Lam', which are able to optimize the sterol composition of cell membranes rapidly under conditions of stress. As in humans, a Niemann-Pick protein NCR1 integrates sterols into the lysosomal membrane prior to further distribution as part of the mechanism of sterol homeostasis. Some antifungal drugs are targeted against ergosterol, either by binding to it to cause damaging cellular leakage, or to prevent its synthesis from lanosterol. Many mutants defective in ergosterol biosynthesis have been isolated, and these have yielded a great deal of information on the features of the sterol molecule required for its structural role in membranes of yeast and fungi. Ergosterol stabilizes the liquid-ordered phase in the same manner as cholesterol and also forms raft microdomains with sphingolipids in membranes, whereas lanosterol does not. It is also evident that ergosterol has a multiplicity of functions in the regulation of yeast growth. Under some conditions, especially those that retard growth, a high proportion of the sterols in yeasts can be in esterified form, where they are stored in lipid droplets. Ergosterol esters are synthesized in yeast by enzymes (ARE1 and ARE2), which are related to ACAT-1 and ACAT-2 that perform this function in animals, and both transfer an activated fatty acid to the hydroxyl group at the C3-position of a sterol molecule. In addition, specific sterol ester hydrolases that catalyze the reverse reaction have been characterized from yeasts, two in lipid droplets and one at the plasma membrane. Many fungal species and slime molds contain steryl glycosides (ergosteryl β-monoglucopyranosides in the former), but they are present at very low levels only in the widely studied yeast Saccharomyces cerevisiae. Most fungi conjugate the 3β-hydroxyl group of ergosterol with aspartate in an RNA-dependent reaction catalyzed by an ergosteryl-3β-O-L-aspartate synthase, with the reverse reaction using a dedicated hydrolase. A phylogenomic study has shown that this pathway is conserved across higher fungi (except S. cerevisiae), including pathogens, and it has been suggested that these reactions constitute a homeostasis system with a potential impact upon membrane remodeling, trafficking, antimicrobial resistance, and pathogenicity. Bacterial Sterols Hopanoids take the place of sterols in many species of bacteria, but it has long been recognized that some bacteria take up cholesterol and other sterols from host animals for use as membrane constituents. Indeed, an external source of sterols is required for growth in species of Mycoplasma. In addition, there have been a number of reports of the biosynthesis of sterols by various bacterial species, although a high proportion of these appears now to have been discounted because of fungal contamination. In particular, the possibility of sterol biosynthesis in cyanobacteria has been controversial, and molecular biology studies have yet to detect the presence of the required enzyme squalene epoxide cyclase. That said, there is good evidence that a few species of prokaryotes at least have the capacity to synthesize sterols de novo. Among the eubacteria, certain methylotrophs (Methylobacterium and Methylosphaera species) produce mono- and dimethyl sterols, including lanosterol. Similarly, some soil bacteria produce 4‑desmethylsterols. It has now been established from gene sequence studies that a few bacteria contain enzymes of the sterol biosynthesis pathway such as oxidosqualene cyclase, but as these have no obvious evolutionary link it seems probable that they were acquired via lateral transfer from eukaryotes. Plant Sterols in the Human Diet The absorption of dietary plant sterols and stanols in humans is low (0.02-3.5%) compared to cholesterol (35-70%), although there are similar amounts in an average Western diet. The explanation is believed to be that the Niemann-Pick C1-like protein 1 (NPC1L1), which is responsible for cholesterol absorption in enterocytes does not take up plant sterols efficiently, while two transporters (ABCG5 and ABCG8) redirect any that are absorbed back into the intestinal lumen. In some rare cases, increased levels of plant sterols in plasma serve as markers for an inherited lipid storage disease (phytosterolemia) caused by mutations in the enterocyte transporters. Among many symptoms, accelerated atherosclerosis is often reported although the reasons for this are not clear. There is evidence that while plant sterols can substitute for cholesterol in maintaining membrane function in mammalian cells, they can exert harmful effects by disrupting cholesterol homeostasis. This may be relevant to the brain especially, since phytosterols are able to cross the blood-brain barrier, although they cannot be oxidized enzymatically because of the alkyl moiety on C24. In contrast, dietary supplements of plant sterols have been reported to have anti-cancer effects. Substantial amounts of phytosterols are available as by-products of the refining of vegetable oils and of tall oil from the wood pulp industry. As it appears that they can inhibit the uptake of cholesterol from the diet and thereby reduce the levels of this in the plasma low-density lipoproteins, there is an increasing interest in such commercial sources of plant sterols to be added as "nutraceuticals" to margarines and other foods, Hydrogenated phytosterols or "stanols" are also used for this purpose, and studies suggest they are as effective as sterols in reducing LDL cholesterol. The consensus amongst experts in the field (including the FDA in the USA) is that such dietary supplements do indeed have the effects claimed and such claims can be used in advertising of commercial products, with the important caveat that there are no randomized, controlled clinical trial data that establish ensuing benefits to health, especially with respect to cardiovascular disease. Other pharmacological effects are under investigation, and there may be beneficial effects for the development of the human fetus and newborn, and for the treatment of non-alcoholic steatohepatitis, inflammatory bowel diseases ,and allergic asthma. It is not clear whether oxy-phytosterols are generated in animal tissues, but those produced by enzymatic or non-enzymatic means can enter the food chain, especially when they are produced during cooking. Although they are not efficiently absorbed, 7-keto-sitosterol and 7-keto-campesterol have been detected in human plasma and have the potential to exert a variety of biological effects. For example, they have pro-atherogenic and pro-inflammatory properties in animal models.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/21%3A_Lipid_Biosynthesis/21.05%3A_Biosynthesis_of_Cholesterol_and_Steroids.txt

Search Fundamentals of Biochemistry By William (Bill) W. Christie and Henry Jakubowski. This section is an abbreviated and modified version of material from the Lipid Web, an introduction to the chemistry and biochemistry of individual lipid classes, written by William Christie. Isoprenoids: 1. Tocopherols and Tocotrienols (Vitamin E) Tocopherols and tocotrienols constitute a series of related benzopyranols (or methyltocols) that are synthesized in plants and other photosynthetic organisms, where they have many important functions but especially as part of a complex web of antioxidants that protect plants from the activities of reactive oxygen species (ROS). First described in 1922 as a dietary factor essential to prevent fetal reabsorption in rats, it was soon understood that plants contained a fat-soluble vitamin (vitamin E) that is essential for innumerable aspects of animal development. Of these many related molecules, only one form, i.e., α‑tocopherol, is recognized as having vitamin E activity in humans in that it prevents a spectrum of human deficiency diseases termed 'ataxia', which is characterized by very low concentrations of α‑tocopherol in plasma. The progression of the disease can be prevented by the administration of α-tocopherol only, although the pathogenic mechanism appears to be uncertain. While all tocopherols are known to be powerful lipid-soluble antioxidants in vitro at least, α‑tocopherol has indirect roles in signal transduction and gene expression in animal tissues. On the other hand, specific functions (non-vitamin E) for other tocopherol forms and their metabolites in animal tissues are now being revealed. Vegetable oils are a major dietary source of vitamin E for humans. Structure and Occurrence In the tocopherols, the C16 side chain is saturated, and in the tocotrienols it contains three trans double bonds. Together, these two groups are termed the tocochromanols. In essence, the tocopherols have a 20-carbon phytyl tail (including the pyranol ring) and four chiral centers in total, with variable numbers of methyl groups attached to the benzene ring, while the tocotrienols have a 20-carbon geranylgeranyl tail with double bonds at the 3', 7' and 11' positions relative to the ring system. Tocopherols contain three chiral carbons, one at C2 in the chromanol ring and two in the side chain at C4′ and C8′ with R,R,R stereochemistry. The four main constituents of the two classes are termed - alpha (5,7,8-trimethyl), beta (5,8-dimethyl), gamma (7,8-dimethyl) and delta (8-methyl). In contrast to the tocopherols, the tocotrienols have only one chiral center. Plastochromanol-8 is an analog of γ-tocotrienol with a much longer side-chain. Their structures are shown in Figure \(1\). The tocochromanols are only synthesized by plants and other oxygenic photosynthetic organisms, such as algae and some cyanobacteria, but they are essential components of the diet of animals. Of these, only natural R,R,R-α-tocopherol is now designated ‘vitamin E’, as explained below, although the other tocopherols are sometimes termed ‘vitamers’ (some claim incorrectly - nor should all forms be termed isomers strictly speaking). In the USA, the current recommended dietary allowance (RDA) is 15 mg α‑tocopherol daily for adults. In plants, there is a great range of tocochromanol contents and compositions, and photosynthetic plant tissues contain from 10 to 50 μg tocochromanols per g fresh weight. α‑Tocopherol only is present in photosynthetic membranes of plant leaves, while γ-tocopherol and other forms are found principally in fruits, seeds, and nuts. While tocopherols are present in all photosynthetic organisms, the tocotrienols are found only in certain plant families. Seed oils are a major source for the human diet and the compositions of tocopherols in some unrefined oils are listed in Table 1. Sunflower and olive oils are good sources of α‑tocopherol and palm oil of the tocotrienols. In general, tocotrienols tend to be abundant only in seeds and fruits, especially of monocots such as wheat, rice, and barley, though a major commercial source is palm oil. In leaf tissue, α-tocopherol is often the main form, while γ-tocopherol is the primary tocopherol of many seeds. Plastochromanol-8 was first found in leaves of the rubber tree (Hevea brasiliensis) but has since been found in many other plants including rapeseed and maize, but usually at lower levels than of the tocopherols. In addition, tocopherol esters of fatty acids occur in plant tissues, where they may be an inert storage form, but unesterified tocopherols are not released during digestion in animals so they may not make a contribution to vitamin E activity. Table \(1\): Tocopherol and tocotrienol contents (mg/Kg) in some seed oils. α-T* β-T γ-T δ-T α-TT* β-TT γ-TT δ-TT palm 89 - 18 - 128 - 323 72 soybean 100 8 1021 421 - - - - maize 282 54 1034 54 49 8 161 6 sunflower 670 27 11 1 - - - - rapeseed 202 65 490 9 - - - - * Abbreviations: T, tocopherol; TT, tocotrienol Data from: Gunstone, F.D., Harwood, J.L. and Padley, F.B. The Lipid Handbook (Second Edition) (Chapman & Hall, London)(1994). An unusual tocopherol that has been termed marine-derived α-tocomonoenol is found together with α-tocopherol in a wide range of marine fish species, where it appears to be a more efficient scavenger of free radicals at low temperatures. A related isomer with a Δ11 double bond has been found in palm oil and kiwi fruit. While pumpkin seeds contain both α- and γ-tocomonoenols, other plant species contain β, γ- and δ-tocomonoenols with unsaturation in the terminal isoprene unit of the side chain. Tocochromenols or 3,4-dehydrotocopherols, i.e., with a double bond in the pyranol ring, are also known in addition to more complex tocopherol-like molecules. The structures of tocomonoenols are shown in Figure \(2\). α-Tocopherol is a minor but ubiquitous component of the lipid constituents of animal cell membranes (non-raft domains), with estimates ranging from one molecule of tocopherol to from 100 to 1000 molecules of phospholipids, depending on the membrane. The hydrophobic tail lies within the membrane, as might be expected, and the polar head group is orientated towards the surface but below the level of the phosphate moieties of the phospholipids. There may be some limited hydrogen bonding between the hydroxyl groups and phosphate depending on the degree of hydration of the membrane. On the other hand, there is a strong affinity of α-tocopherol for polyunsaturated fatty acids, where the chromanol unit may interact with the double bonds, suggesting that tocopherol is located deep within the membrane. α-Tocopheryl phosphate has recently been detected at low levels in plasma, liver, and adipose tissue. Its structure is shown in Figure \(3\). Together with catabolic tocopherol metabolites, it has important biological properties. During the refining of vegetable oils, much of the natural tocopherols is lost or destroyed. Most commercial vitamin E is therefore prepared by chemical synthesis with trimethylhydroquinone and phytyl bromide as the precursors. The resulting product is a mixture of eight stereoisomers (from R,R,R- to S,S,S-methyl groups) of α-tocopherol, with the various stereoisomers differing by a factor of two in biologic activity, as a consequence of the stereochemistry of position 2 in the chromanol ring (i.e., 2S-α- compared to 2R-α-tocopherol). It is usually administered as the acetate derivative in vivo. Tocopherols are not usually regarded as effective antioxidants in the polyunsaturated seed oils of commerce, and at higher concentrations can even act as pro-oxidants, although the reasons for this are not understood. Biosynthesis and Functions of Tocochromanols in Plants The mechanism of biosynthesis of tocopherols has been elucidated and involves the coupling of phytyl diphosphate with homogentisic acid (2,5‑dihydroxyphenylacetic acid), followed by cyclization and methylation reactions. The plant chloroplast is the site of biosynthesis, and most of the enzymes are located on the inner membrane of the chloroplast envelope, although there is increasing evidence that plastoglobules associated with the thylakoid membrane may be involved. The pathway for biosynthesis of tocopherols is shown in Figure \(4\). The aromatic amino acid tyrosine can be considered the basic precursor, and this is oxidized to p-hydroxypyruvic acid, which in the first committed step is converted to homogentisic acid by the enzyme p-hydroxyphenylpyruvate dioxygenase. Homogentisic acid is condensed with phytyl diphosphate, derived from phytol obtained from hydrolysis of chlorophyll, in a reaction catalyzed by a prenyl transferase to yield 2-methyl-6-phytyl-plastoquinol, which is first methylated to form 2,3-dimethyl-5-phytyl-1,4-benzoquinol and then converted by the enzyme tocopherol cyclase to γ-tocopherol. A further methylation reaction produces α-tocopherol, while modifications to the pathway produce β- and δ-tocopherols, together with plastoquinones and thence plastochromanol-8. Tocotrienols and tocomonoenols result from a similar series of reactions but with geranylgeranyl diphosphate and tetrahydro-geranylgeraniol diphosphate, respectively, as substrates in the condensation step. The isoprenoid precursors are synthesized in the plastid also by the non-mevalonate or 'MEP' pathway. In plants, tocopherols are found mainly in the chloroplasts of green tissues, but they are also present in seeds, fruits, roots and tubers. They are especially important as antioxidant molecules, limiting the damage from photosynthesis-derived reactive oxygen species during conditions of oxidative stress, including high-intensity light stress, and the mechanisms for this antioxidant activity are discussed below. However, recent studies seem to suggest that they are just one of a number of different components that are involved in photoprotection. Certainly, any tocochromanol peroxy radicals formed must be converted back to the original compounds by the concerted action of other plant antioxidants, for example by ascorbate, glutathione, ubiquinol or lipoic acid, and antioxidant enzymes, including superoxide dismutase, catalase, and peroxidases. Tocopherols are essential for the control of non-enzymatic lipid peroxidation during seed dormancy and germination of seedlings. In their absence, elevated levels of malondialdehyde and phytoprostanes are formed, and there can be inappropriate activation of plant defense responses. There is evidence that tocopherols play a part in intracellular signaling in plants in that they regulate the amounts of jasmonic acid in leaves, via modulating the extent of lipid peroxidation and gene expression, and so influence plant development and stress responses. Thus, by controlling the degree of lipid peroxidation in chloroplasts (redox regulation), they limit the accumulation of lipid hydroperoxides required for the synthesis of jasmonic acid, which in turn regulates the expression of genes that affect a number of abiotic stress conditions, including drought, salinity and extremes of temperature. The translocation of enzymes to the plasma membrane is regulated by tocopherols, possibly by modulating protein-membrane, altering membrane microdomains (lipid rafts), or by competing for common binding sites within lipid transport proteins. In addition, tocopherols are required for the development of the cell walls in phloem transfer cells under cold conditions. It appears that α- and γ‑tocopherol and the tocotrienols may each have distinct functions. For example, γ-tocopherol is reportedly more potent than α-tocopherol in protecting plants from the harmful effects of osmotic stresses and is important for the longevity of seeds. Efforts are underway to increase the tocopherol levels in plants by selective breeding and genetic manipulation with the aim of producing crops with greater potential health benefits to consumers and perhaps for the plants per se. Tocopherols Metabolism in Animals In animals, the first step in the digestion of tocopherols is their dissolution with other lipids in mixed micelles in the intestines. All tocopherol forms are absorbed to a similar extent in the intestines by means of transporters in the enterocyte apical membrane that have a broad specificity for hydrophobic molecules, such as cholesterol, vitamin D, and carotenoids. These include scavenger receptor class B type I (SR-BI), the CD36 protein, and NPC1-like intracellular cholesterol transporter 1 (NPC1L1). However, some passive diffusion cannot be ruled out. Transport across the enterocyte may involve cytoplasmic transporters or clathrin-coated vesicles before the tocopherols are incorporated into chylomicrons in free form in the Golgi apparatus for release into the lymph. At the liver, α-tocopherol specifically is taken up from the chylomicrons by a receptor-mediated mechanism with the aid of a specific tocopherol-binding protein (the α-tocopherol transfer protein (α-TTP)), i.e., a 30,500 Da cytosolic protein that has a marked affinity for α-tocopherol and can enhance its transfer between membranes. This recognizes α-tocopherol by the three methyl groups and hydroxyl on the chromanol ring and by the structure and orientation of the phytyl side chain. It is the chief regulator of whole body α-tocopherol status and is expressed primarily in the cytosol of hepatocytes in the liver, but has been reported (in much lower concentrations) in other tissues, such as the placenta. α-TTP ultimately regulates the egress of α-tocopherol selectively from hepatocytes with the aid of the ATP-binding cassette proteins ABCA1 and ABCG for conveyance in the plasma lipoproteins, mainly in the very-low-density lipoproteins or VLDL (and thence to LDL) and HDL in humans, to the peripheral tissues (together with much smaller amounts of γ-tocopherol). Most of the other tocopherol forms are directed toward catabolism. Once in the circulation, tocopherol can exchange spontaneously between membranes and lipoproteins, and no specific transport protein for vitamin E in plasma has yet been described. Transfer of tocopherols from the VLDL to peripheral tissues occurs as triacylglycerols are hydrolyzed by the enzyme lipoprotein lipase, while that in LDL is processed via the LDL receptor-mediated uptake pathway. Within cells of peripheral tissues, including the central nervous system, α-TTP functions in transporting α-tocopherol to wherever it is required in membranes, a process that appears to be aided by phosphatidylinositol metabolites. In the brain, tocopherol is transported by apo-E rich lipoproteins. Concentrations of tocopherols can vary appreciably amongst tissues, with most in adipose tissue and adrenals, less in kidney, heart, and liver, and least in the erythrocytes. The "α-tocopherol salvage pathway" is partly due to this process and partly to selective oxidation (see below), and the result is a 20- to 30-fold enrichment of α‑tocopherol in plasma (average concentration 22-28 μM) relative to the other tocopherols. Thus, the process of conservation of one specific tocopherol appears to determine the relative vitamin E activities of the tocopherols and tocotrienols in vivo, rather than their individual potencies as antioxidants as measured in model systems in vitro. Only α‑tocopherol (including synthetic material) or natural mixtures containing this can be sold under the label 'Vitamin E'. γ‑Tocopherol is the second most abundant form in plasma, and it is present in relatively greater proportions in the skin, adipose tissue, and skeletal muscle, where it has some specific biological properties that are distinct from those of α-tocopherol. Although tocotrienols are more potent antioxidants in vitro, they are not usually detected in tissues, although they are believed to have some important functions. Catabolism: The unwanted surplus of tocochromanols other than α-tocopherol may be excreted in the urine and feces in the form of carboxy-chromanols, including the so-called 'Simon metabolites' - tocopheronic acids (carboxyethylhydroxychromans, CEHC) and tocopheronolactones, after oxidative cleavage of much of the phytyl tail, although these are normally detected in the form of conjugates as sulfate or glucuronidate esters, the forms in which they are excreted in feces and urine. For example for illustrative purposes in liver cells, the first step in the catabolism of γ-tocopherol is ω-hydroxylation by cytochrome P450 (CYP4F2) at the 13' carbon to form γ-13'-hydroxychromanol in the endoplasmic reticulum, followed by ω-oxidation in the peroxisomes to produce γ‑13'‑carboxychromanol, and finally by stepwise β‑oxidation in the mitochondria to cut off two or three carbon moieties from the phytyl chain in each cycle. These steps are shown in Figure \(5\). Various carboxychromanol intermediates have been identified for all of the tocopherols together with forms in which the hydroxyl group is sulfated in human cell cultures in vitro; sulfated carboxychromanols are the main tocopherol metabolites in the plasma of rodents. As the vitamin E ω-hydroxylase has a high affinity for the tocopherols other than the α-form and does not attack that bound to the α-tocopherol transfer protein, this provides a further specific enhancement of the α‑tocopherol concentration in plasma relative to the others. Some of these catabolic metabolites may have some biological activity in their own right. For example, carboxyethylhydroxychromans derived from γ-tocopherol were reported to induce apoptosis in cancer cells and to have anti-inflammatory effects by inhibition of cyclooxygenases and 5-lipoxygenase (see below). Tocotrienols are catabolized in a similar manner, but with additional steps in which the double bonds are reduced prior to oxidation; the final carboxyethylchromanols are the same as for tocopherols. Tocopherols as Antioxidants Although the syndrome associated with a lack of vitamin E in the diet of animals has been known for decades, the mode of action and specific location of tocopherols in cell membranes are not clearly understood. Several theories have been proposed to explain the functions of vitamin E in animal cells. From studies in vitro, it has long been believed that a major task is to act as an antioxidant to inhibit, decrease, delay, or prevent oxidative damage to unsaturated lipids or other membrane constituents and thence to tissues by scavenging free radicals. For example, vitamin E administration can prevent lipid peroxidation and hepatotoxicity upon exposure to the free radical-generating agent carbon tetrachloride. Lipid peroxidation is also a cause of ferroptosis, an iron-dependent form of nonapoptotic cell death. However, tocopherols have functions other than as antioxidants. In non-biological systems such as foods, cosmetics, pharmaceutical preparations, etc., tocopherols are invaluable as antioxidant additives. Because of their lipophilic character, tocopherols are located in the membranes or with storage lipids where they may be available immediately to interact with lipid hydroperoxides, such as those described in more detail in our web pages on isoprostanes, reactive aldehydes, and oxidized phospholipids. In brief, Reactive Oxygen Species (ROS), of which innumerable forms, exist can be derived by enzymatic or non-enzymatic means and produce superoxide anions and other peroxyl radicals. Superoxide radicals (O2•-) ultimately generate highly toxic hydroxyl (OH) or alkoxyl radicals, which can abstract a hydrogen atom from bis-allylic methylene groups of polyunsaturated fatty acids under aerobic conditions in vivo in animals and plants to generate lipid peroxyl radicals (LOO) and hydroperoxy-fatty acids. Singlet oxygen (1O2 or O=O) is an especially important ROS (non-radical) in photosynthetic tissues of plants. As radical generation is not enzymatic, all methylene groups between two cis double bonds can potentially be involved in the reaction, although not necessarily to the same degree. Tocopherols react rapidly in a non-enzymic manner unlike many other cellular antioxidants, which are dependent on enzymes, to scavenge lipid peroxyl radicals, i.e., the chain-carrying species that propagate lipid peroxidation. In model systems in vitro, all the tocopherols (α > γ > β > δ) and tocotrienols are good antioxidants, with the tocotrienols being the most potent. In general, the oxidation of lipids is known to proceed by a chain process mediated by such free radicals, in which the lipid peroxyl radical serves as a chain carrier. In the initial step of chain propagation, a hydrogen atom is abstracted from the target lipid by the peroxyl radical as shown in Figure \(6\). The main function of α-tocopherol is to scavenge the lipid peroxyl radical before it is able to react with the lipid substrate as shown in Figure \(7\). The potency of an antioxidant is determined by the relative rates of reactions (1) and (2). When a tocopheroxyl radical is formed, it is stabilized by the delocalization of the unpaired electron about the fully substituted chromanol ring system rendering it relatively unreactive, thus preventing propagation of the chain reaction. This also explains the high first-order rate constant for hydrogen transfer from α-tocopherol to peroxyl radicals, as studies of the relative rates of chain propagation to chain inhibition by α-tocopherol in model systems have demonstrated that α-tocopherol is able to scavenge peroxyl radicals much more rapidly than the peroxyl radical can react with a lipid substrate. In biological systems, oxidant radicals can spring from a number of sources, including singlet oxygen, alkoxyl radicals, superoxide, peroxynitrite, nitrogen dioxide, and ozone. α-Tocopherol is most efficient at providing protection against peroxyl radicals in a membrane environment. The reaction of the tocopheroxyl radical with a lipid peroxyl radical, as illustrated, yields 8α-substituted tocopherones, which are readily hydrolyzed to 8α-hydroxy tocopherones that rearrange spontaneously to form α-tocopherol quinones. In an alternative pathway, the tocopheroxyl radical reacts with the lipid peroxyl radical to form epoxy-8α-hydroperoxytocopherones, which hydrolyze and rearrange to epoxyquinones. Tocopherol dimers and trimers may also be formed as minor products. These reactions are shown in Figure \(8\). Free radical-mediated lipid peroxidation is the major pathway of lipid oxidation taking place in humans, and α-tocopherol is a major antioxidant, but it does not scavenge the nitrogen dioxide radical, carbonate anion radical, and hypochlorite efficiently. Vitamin E forms with an unsubstituted 5-position, such as γ-tocopherol, are an exception to the rule that the various tocopherols have similar antioxidant properties in that they are able to trap electrophiles, including Reactive Nitrogen Species (RNS), which are enhanced during inflammation. The enzyme nitric oxide synthase is capable of continuously producing a large amount of nitric oxide (NO), which can react with superoxide to produce peroxynitrite (ONOO-), a potent and versatile oxidant that can attack a wide range of biological targets. It induces lipid peroxidation and nitrates aromatic compounds and unsaturated fatty acids while isomerizing cis-double bonds in fatty acids to the trans-configuration. γ-Tocopherol is superior to α‑tocopherol in detoxifying the NO2 radical and peroxynitrite with formation of 5-nitro-γ-tocopherol, as shown in Figure \(9\). Figure \(\PageIndex{xx}\): This occurs in vivo, and the concentrations of 5-nitro-γ-tocopherol have been shown to be elevated in the plasma of subjects with coronary heart disease and in carotid-artery atherosclerotic plaque. In plant and animal tissues, tocopherols can be regenerated from the tocopheroxyl radicals in a redox cycle mediated by a number of endogenous antioxidants, including vitamins A and C and coenzyme Q, and this must greatly extend their biological potency. Vitamin C (ascorbic acid) may be especially important in aqueous systems, although it may also act at the surface of membranes, to regenerate α-tocopherol, while in turn being oxidized to dehydroascorbic acid. This can be regenerated to the reduced form by glutathione (GSH) with the production of glutathione disulfide (GSSG), which can subsequently be enzymatically reduced by glutathione reductase with NAD(P)H as a cofactor. In plants, an NAD(P)H-dependent quinone oxidoreductase is involved at an early stage of the regeneration process, while tocopherol cyclase, an enzyme involved in the biosynthesis of tocopherols, re-introduces the chromanol ring. These linked cycles (the antioxidant network) are shown in Figure \(10\). Thus, tocopherols are only one component of a complex web of metabolites and enzymes in tissues that have antioxidant activities and act by various mechanisms, including the stimulation of genes involved in signaling responses to environmental stresses. One antioxidant mechanism involves the removal of free radicals and reactive species by enzymes such as superoxide dismutase, catalase, and glutathione peroxidase, while electron donors, such as glutathione, tocopherols, ascorbic acid, vitamin K, coenzyme Q, and thioredoxin, scavenge free radicals also. Metal-binding proteins such as transferrin, metallothionein, haptoglobin, and ceruloplasmin have antioxidant activity by sequestering pro-oxidant metal ions, such as iron and copper, although some metals such as selenium and zinc are in fact antioxidants. Other antioxidants, including flavonoids, carotenoids, and phenolic acids in addition to tocopherols, enter animal tissues via the food chain. Although the discussion here has been limited to the effects upon lipids, free radicals can cause damage to proteins, DNA, and indeed virtually any native substance in living organisms. Biological Functions of Tocochromanols in Animals Vitamin E deficiency has been detected in patients with fat malabsorption, cystic fibrosis, Crohn's disease, liver disease, and pancreatic insufficiency, and in premature infants. Impairment of the normal functions of the immune system has been demonstrated in animals and humans in vitamin E deficiency, and this can be corrected by vitamin E repletion. It also displays activity against nonalcoholic hepatosteatosis. Although there are various proposals for the pathogenic mechanism, none as yet appears to be generally accepted. After the discovery of the effects of vitamin E on fertility in studies with laboratory animals, its importance was documented for the development of tissues and organs such as brain and nerves, muscle and bones, skin, bone marrow, and blood, most of which are specific to α-tocopherol. However, there is no evidence for an effect of vitamin E on fertility in humans, as was originally found in the rat. The rare genetic disorder “Ataxia with Isolated Vitamin E Deficiency” or “AVED” is the result of mutations in the gene coding for α-TTP. It is caused by the death of cerebellar Purkinje cells, but administration of α-tocopherol prevents this and the subsequent development of clinical symptoms of the disease. There appears to be little doubt that tocopherols inhibit many of the enzymes associated with inflammation in vitro in animals, and may contribute to the amelioration and treatment of some chronic diseases. However, it has been argued that data on the effects of vitamin E on biomarkers of oxidative stress in vivo are inconsistent. Oxidized metabolites of vitamin E, i.e., that have reacted as antioxidants, are barely detectable in tissues, and vitamin E maintenance in vivo does not appear to have been clearly associated with its regeneration. There appear to be significant differences between results obtained in studies with laboratory animals in comparison to those in humans. Thus, suggestions that dietary supplements of vitamin E may reduce the rate of oxidation of lipids in low-density lipoproteins in humans and thence the incidence or severity of atherosclerosis have not been confirmed by clinical intervention studies, although benefits in some conditions have been claimed. Indeed, there are suggestions that excessive vitamin E supplementation may even be harmful. One study has suggested that relatively high doses of natural α-tocopherol over a long period are required to demonstrate a significant reduction in the levels in the urine of F2 isoprostanes, which are considered to be the most reliable marker for oxidative stress in vivo. While there are many fat-soluble antioxidants in the diet, only α-tocopherol is a vitamin. It has even been suggested that tocopherol may be protected from functioning as an antioxidant in some tissues in vivo through a network of cellular antioxidant defenses, such that tocopherols are utilized only when other antioxidants are exhausted, although there is no experimental proof of this hypothesis. At the cellular level, RRR-α-tocopherol has been shown to inhibit protein kinase C, and in the process, it inhibits the assembly and radical-producing activity of NADPH oxidase in monocytes. Similarly, vitamin E suppresses the expression of xanthine oxidase, a source of reactive oxygen species, in the liver. It is thus possible that α-tocopherol is able to diminish the levels of free radicals by preventing their production and not by scavenging them. Its physical presence in membranes adjacent to polyunsaturated fatty acids may thus limit autoxidation. With the discovery that the antioxidant effects of various tocopherols and tocotrienols have little relation to their vitamin E activities in vivo has come the realization that they have other functions in tissues, most of which are specific to α-tocopherol. Most current research is concerned with how vitamin E and its metabolites act in signaling and controversially in the regulation of gene activity. While it is certainly true that most other vitamins are essential cofactors for specific enzymes or transcription factors, no receptor that binds specifically to vitamin E has yet been discovered. By preventing the increase of peroxidized lipids that alter both metabolic pathways and gene expression profiles within tissues and cells, it may act indirectly as a regulator of genes connected with tocopherol catabolism, lipid uptake, collagen synthesis, cellular adhesion, inflammation, the immune response and cell signaling. Vitamin E affects a number of transcription factors in this manner, including peroxisome proliferator-activated receptor gamma (PPARγ), nuclear factor erythroid-derived 2 (NRF2), nuclear factor kappa B (NFκB), RAR-related orphan receptor alpha (RORα), estrogen receptor beta (ERβ), and the pregnane X receptor (PXR). α-Tocopherol and its metabolites are believed to modulate the activity of several enzymes involved in signal transduction, including protein kinases and phosphatases, lipid kinases and phosphatases, and other enzymes involved in lipid metabolism, but especially those with inflammatory properties such as lipoxygenases, cyclooxygenase-2, and phospholipase A2. While the credentials of tocopherols as antioxidants in vivo have been doubted, this does not preclude a role in the inhibition of oxidative enzymes, especially in relation to the function of the immune system. For example, vitamin E regulates T cell function directly by its effects upon T cell membrane integrity, signal transduction, and cell division, and it also functions indirectly by affecting eicosanoids and related inflammatory mediators generated from other immune cells. Various tocopherols and tocotrienols have been shown to suppress COX-2 involvement in prostaglandin (PGD2 and PGE2) synthesis in lipopolysaccharide-activated macrophages. In addition, it has been established that the 13'-carboxy metabolite of α-tocopherol (α-T-13'-COOH) and other tocopherol ω-carboxylates are potent allosteric inhibitors of 5-lipoxygenase, a key enzyme in the biosynthesis of the inflammatory leukotrienes. α-T-13'-COOH accumulates in immune cells and inflamed exudates both in vitro and in vivo in mice, and it has even been suggested that the immune regulatory and anti-inflammatory functions of α-tocopherol depend on this endogenous metabolite. The structure of α-T-13'-COOH is shown in Figure \(11\). α-Tocopherol has a stimulatory effect on the dephosphorylation enzyme, protein phosphatase 2A, which cleaves phosphate groups from protein kinase C, leading to its deactivation. The mechanism may involve the binding of vitamin E directly to enzymes in order to compete with their substrates, or it may change their activities by redox regulation. It may also compete for common binding sites within lipid transport proteins, and so may alter the traffic of lipid mediators indirectly with effects upon their signaling functions and enzymatic metabolism. For example, it binds to albumin as well as to a specific α-tocopherol-associated protein (TAP), and in the latter form especially it inhibits the phosphoinositide 3-kinase. It has been suggested that vitamin E may have a secondary role in stabilizing the structure of membranes, or it may interact with enzymes in membranes to interfere with binding to specific membrane lipids, or it may affect membrane microdomains such as lipid rafts. Evidence suggests that the biological activities of β-, γ- and δ-tocopherols do not reflect their behavior as chemical antioxidants, but anti-inflammatory, antineoplastic, and natriuretic actions have been reported. Some non-antioxidant effects of γ-tocopherol in tissues in relation to reactive nitrogen oxide species have been observed, but the specificity of these in vivo is not yet certain. In addition, anti-inflammatory properties have been described that have been attributed to a chain-shortened metabolite. Beneficial effects against cancer cells in vitro have been observed that have been ascribed to scavenging of reactive nitrogen species, since such effects are not seen with α-tocopherol. On the other hand, vitamin E and its derivatives are believed to regulate tumor cells by activating the mitogen-activated protein kinase (MAPK) signaling pathway. Tocotrienols have been shown to have neuroprotective effects and to inhibit cholesterol synthesis. They reduce the growth of breast cancer cells in vitro, possibly by influencing gene expression by interaction with the estrogen receptor-β. When administered in combination with either standard antitumor agents as in chemotherapy or with natural compounds with anticancer activity, they are reported to exert a synergistic antitumor effect on cancer cells. γ-Tocotrienol is reported to be an inducer of apoptosis via endoplasmic reticulum stress, while α-tocotrienol may be neuroprotective by inhibition of lipoxygenase activity. Although anti-obesity and anti-diabetic effects have been observed in mice, clinical trials with humans appear to have given inconclusive results. These properties are largely distinct from those of the tocopherols, and the pharmaceutical potential of tocotrienols against cancer, bone resorption, diabetes, and skin, cardiovascular and neurological diseases are currently being studied. The biological functions of α-tocopheryl phosphate are slowly being revealed. In addition to being a possible storage or a transport (water-soluble) form of tocopherol, it is involved in cellular signaling and regulates a number of genes, including those involved in angiogenesis and vasculogenesis, in a different manner from α‑tocopherol per se. As it lacks the free hydroxyl group, it cannot act directly as an antioxidant, and some consider it to be the biologically active form of the vitamin. It is certainly more active in a number of biological systems in vitro than α-tocopherol, so these effects cannot be ascribed to the hydrolyzed molecule, and in some instances, it is antagonistic to α-tocopherol, for example in its activity towards phosphatidylinositol 3-kinase. On the other hand, activation requires a kinase, while a phosphatase is needed to make the system reversible, but neither has yet been identified. Synthetic phosphate derivatives of γ-tocopherol and α-tocopheryl succinate are known to have potent anti-cancer properties. Isoprenoids: 2. Retinoids (Vitamin A) That a dietary factor was involved in visual acuity was known to the ancient Egyptians and Greeks, but it was the 1930s before the importance of the carotenoids and their metabolites was recognized, and β-carotene and retinol were fully characterized. It is now recognized that vitamin A activity now resides in the metabolites retinol, retinal and retinoic acid, and in several provitamin A carotenoids, most notably β-carotene. A share in the Nobel Prize for Medicine in 1967 was awarded to George Wald, who over many years showed how retinol derivatives (named for their function in the retina) constituted the chemical basis of vision. Now, it is recognized that retinol, retinoic acid, and their many metabolites have innumerable other functions in human metabolism from embryogenesis to adulthood, including growth and development, reproduction, cancer, and resistance to infection. They are important natural antioxidants with benefits to health, although some potentially harmful properties have been reported. Carotenoids are a class of highly unsaturated terpenoids that occur in innumerable molecular forms (>1000). They are common colorful pigments of plants, fungi, and bacteria, of vital importance to photosynthesis, and as dietary constituents, they can add ornament to some animal species. Apart from acting as precursors of retinoids, carotenoids per se appear to have a relatively limited range of functions in animal tissues, but they are important to vision and as antioxidants, especially in the skin. They do of course have important functions in plants and lower organisms where they originate, but this topic can only be dealt with briefly here. Other fat-soluble vitamins tocopherols (vitamin E), vitamin K, and vitamin D are discussed elsewhere. Occurrence and Basic Metabolism of Carotenoids and Retinoids The term ‘vitamin A’ is used to denote retinol (or all-trans-retinol, sometimes termed 'vitamin A1'), together with a family of biologically active C20 retinoids derived from this ('vitamers'). The structure of retinol is shown in Figure \(12\). These are only found in animal tissues, where they are essential to innumerable biochemical processes. However, they cannot be synthesized de novo in animals and their biosynthetic precursors are plant carotenoids with a β-ionone ring (provitamin A), C40 tetraterpenes of which β‑carotene is most the efficient; it is an orange-red pigment that occurs in the photosynthetic tissues of plants and in seed oils. In the human diet in the developed world, plant sources tend to be less important than those from dairy products, meat, fish oils, and margarines, which provide vitamin A per se, although carrots and spinach are good sources of the provitamin. In the U.K., for example, all vegetable spreads must be supplemented with the same level of vitamin A (synthetic retinol or β-carotene) as is found in butter. While most research effort has been focused on retinoids, there is increasing interest in the biological activities of intact carotenoids in animal tissues. The biosynthesis of carotenoids in plants via isopentenyl diphosphate and dimethylallyl diphosphate has much in common with that of the plant sterols, but this is too specialized a topic to be treated at length here. They have many important functions in plants, for example during photosynthesis or as precursors of plant hormones, and these are discussed below. Some crop plants with increased carotene levels are available with the aim of preventing vitamin A deficiency in the populations of developing countries, and further efforts are underway. Non-photosynthetic bacteria produce a different range of carotenoids, some with chain lengths other than C40 (C30 to C50). In animals (including humans), dietary carotenoids such as β-carotene are solubilized with other dietary lipids in mixed micelles with the aid of bile acids, and they are absorbed in the intestines in intact form by a process facilitated by specific receptor proteins. Dietary retinol and retinol esters are absorbed similarly in the intestines, but the latter are first hydrolyzed by pancreatic lipase. Conversion to retinoids leading ultimately to retinol esters occurs in the enterocytes, where dietary β-carotene is subjected to oxidative cleavage at its center, the first step of which is catalyzed by a cytosolic enzyme β-carotene-15,15'-oxygenase‑1 (BCO1), which is specific for carotenes with a β-ionone ring, to yield two molecules of all-trans-retinal, which is reversibly reduced by a retinol reductase to retinol. Xanthophyll carotenoids are absorbed without cleavage mainly. These reactions are shown in Figure \(13\). Based on the incorporation of 18O into the products. it appears that the enzyme that introduces the oxygen atom, β-carotene-15,15'-oxygenase‑1 (BCO1), into the cleavage products is a dioxygenase as both atoms of oxygen in dioxygen are incorporated into products. This is in contrast to a possible mechanism in which only one oxygen atom from dioxygen is added, with the other coming from H2O if the enzyme acted as a monooxygenase. Figure \(14\) shows how oxygen could be introduced in the reaction catalyzed by BCO1 through both possible mechanisms. The dioxygenase mechanism on the right best accounts for the incorporation of 18O isotope of oxygen in dioxygen. Figure \(15\) shows an interactive iCn3D model of the apocarotenoid cleavage oxygenase from Synechocystis, a Retinal-Forming Carotenoid Oxygenase (2BIW) Figure \(15\): Retinal-Forming Carotenoid Oxygenase from Synechocystis (2BIW). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov...dttF958wFPAUP7 The enzyme has a bound carotene analog, (3R)-3-hydroxy-8'-apocarotenol. It contains an active site Fe2+ ion at the end of a hydrophobic tunnel. The ion is ligated by 4 histidine side chains. On binding, three of the trans C=C bonds convert to a distorted cis-trans-cis conformation. The middle trans bond is proximal to the Fe2+ which is ligated by dioxygen, the source of the incorporated oxygen atoms on cleavage. Any unchanged β-carotene and newly formed retinol esters in the enterocytes are incorporated into chylomicrons and released into the lymphatic system and thence into the bloodstream, where some is taken up by peripheral tissues before most is absorbed by the liver. Some intact carotene and other carotenoids are transferred to lipoproteins (LDL and HDL) for transport in plasma, with assistance from specific binding and transport proteins, and for example, carotene can be absorbed at the placental barrier and transferred to the fetus for conversion to retinoids that are essential for development. Within the hepatocytes, retinol esters are hydrolyzed in the late endosomes with the release of free retinol into the cytosol, from which it can be released back into the circulation, converted to retinoids or transferred to hepatic stellate cells for storage in lipid droplets, the main body reservoir of vitamin A. In these specialized cells, retinol is esterified to form retinyl palmitate by the transfer of fatty acids from position sn-1 of phosphatidylcholine, mainly via the action of a membrane-bound lecithin:retinol acyltransferase (LRAT) in the endoplasmic reticulum. There are also lesser acyl-CoA dependent pathways, including an acyl CoA:retinol acyltransferase and even the enzyme diacylglycerol acyltransferase 1 (DGAT1); esterification is facilitated by binding to cellular retinol-binding protein type II (CRBP2). Both retinol and retinoic acid are precursors of a number of metabolites (retinoids), which are required for specific purposes in tissues, by enzymatic modification of the functional groups and geometrical isomerization of the polyene chains. In the liver, activation of the retinol pathway involves first mobilization of the ester, followed by hydrolysis by retinol ester hydrolases, which includes carboxylesterase ES-10. Then, the reversible oxidation of retinol to retinal is carried out by one of several enzymes that include dehydrogenases and various cytochrome P450s, before some retinal is oxidized irreversibly to retinoic acid by enzymes with retinal dehydrogenase activity. On-demand conversion of retinol to retinoic acid occurs by the same mechanisms in other tissues, although for vision, retinol esters serve directly as the substrate for the formation of the visual chromophore 11‑cis-retinal (see below). Retinyl-β-D-glucoside, retinyl-β-D-glucuronide, and retinoyl-β-D-glucuronide are naturally occurring and biologically active metabolites of vitamin A, which are found in fish and mammals. Indeed, the last has similar activity to all-trans-retinoic acid without any of the unwanted side effects in some circumstances. Cleavage of β-carotene at double bonds other than that in the center or of a wider range of other carotenoids occurs by the action of a related enzyme β‑carotene-9',10'-dioxygenase (β‑carotene-oxygenase‑2 or BCO2) in mitochondria, which leads to the formation of similar molecules, i.e. β-apocarotenals and β‑apocarotenones of variable chain-length. While these may exert distinctive biological activities in their own right, there is evidence that they can also be metabolized to form retinal. In the aqueous environment within cells, as well as in plasma, retinol, retinal and retinoic acid are bound to retinoid-binding proteins (RBP), which solubilize, protect and in effect detoxify them. These proteins also have a role in facilitating retinoid transport and metabolism; some are present only in certain tissues, and many are specific for particular retinoids and metabolic pathways. To prevent infiltration through the kidneys, retinol, and holo-RBP form an association in blood with a protein termed transthyretin (TTR), which also serves as a thyroid hormone carrier and is essential for secretion. Normally, vitamin A circulates in plasma as a retinol:RBP:TTR complex with a 1:1:1 molar ratio. Unesterified retinol is the main form of the vitamin that is exported from the liver upon demand, and it is transported in the blood in this bound form in VLDL, LDL, and HDL, with some directly from the diet in the chylomicrons and their remnants. Peripheral tissues have specific receptors to take up what they require, probably after hydrolysis of any esters to retinol by means of the enzyme lipoprotein lipase. Then, retinol dissociates from the protein as it forms a complex with a receptor (STRA6) at a target cell and diffuses through the plasma membrane, a process driven by retinol esterification. The RBP-TTR complex does not bind to retinal and retinoic acid, although these do bind to RBP on its own, and most of the low levels of retinoic acid transported in the blood are bound to albumin. Local levels of retinoic acid are the result of an interplay between enzymes of synthesis, binding, and catabolism. For example, within cells retinoic acid binding proteins (CRABP1 and CRABP2) bind to the newly synthesized retinoic acid, increase its rate of metabolism and protect cells from an excess. In skin, 3,4-dehydroretinoids are synthesized from all-trans retinoids by the desaturase Cytochrome P450 27C1 with the assistance of cellular retinol-binding proteins (CRBPs). 3,4-Dehydroretinol, which is sometimes termed vitamin A2, is shown in Figure \(16\). Its derivative 3,4‑dehydroretinal is used as a visual chromophore in many cold-blooded vertebrates including lampreys, fish, amphibians, and some reptiles (see below). Geranylgeranoic acid has structural similarities to retinoic acid and has been termed an acyclic retinoid, although it has no vitamin A activity. It is synthesized in animal tissues from mevalonate, and together with its 2,3‑dihydro metabolite, has potent anticancer properties. Retinol esters: A relatively small proportion of the cellular retinoids is located in membranes in tissues. Rather, retinol esters, mainly retinyl palmitate, are the main storage form of vitamin A, and they occur in many different organs, including adipose tissue and testes, but chiefly in stellate cells of the liver and pancreas. How the retinol is directed specifically to these cells and enters them prior to esterification is not known. Although hepatic stellate cells are much smaller and less abundant than hepatocytes (only 5 to 8% of all liver cells), they are characterized by cytoplasmic lipid droplets that contain 90-95% of the hepatic retinoids (and up to 80% of the body pool) in addition to other non-retinoid lipids; the lecithin:retinol acyltransferase is the only retinol ester synthase in this instance. In addition, specialized cells in the eye store retinoids essential for vision in the form of lipid droplets. When the supply of retinol in the diet is limited, hepatic stores of retinol esters are mobilized as retinol ester hydrolases are activated to maintain constant circulating retinol levels; hormone-sensitive lipase is the most important of these enzymes, although the adipose tissue triacylglycerol lipase and the lysosomal acid lipase are also involved. Catabolism: All-trans-retinoic acid formation is irreversible, so its synthesis and degradation must be tightly regulated. As a first step in catabolism, the excess is cleared by conversion to more polar metabolites through oxidation by various enzymes of the cytochrome P450 family. Secondly, the water-soluble retinoic acid metabolites, including 4-hydroxy-, 4-oxo- and 18-hydroxy-retinoic acids, conjugate with glucuronic acid and then can be rapidly removed from circulation and eliminated from the body via the kidney. Retinoids and Vision The structure of 11-cis-retinal, which we discussed in Chapter 11, is shown in Figure \(17\) Retinoids are essential for vision, and there is now a good appreciation of how this works at the molecular level. In the eye, uptake of retinol from the circulation is mediated by the transmembrane cell-surface STRA6 receptor of the retinal pigment epithelium, a pigmented monolayer of cells located between the photoreceptors and choroid that nourishes retinal visual cells and catalyzes the release of retinol from retinol-binding proteins and transports it to the cytosol. The process by which light is converted to a signal recognized by the brain, sometimes termed the 'retinoid (visual) cycle', requires a two-cell system beginning in the retinal pigment epithelium and continuing in photoreceptor cells, i.e. retinal rod and cone cells in the eye that contain membranous vesicles that serve as light receptors. Roughly half of the proteins in these vesicles consist of the protein conjugate, rhodopsin, which consists of a protein – opsin – with the retinoid 11-cis-retinal. Each step in the visual process requires specific binding or transport proteins, and especially the interphotoreceptor retinoid-binding protein (IRBP). All-trans-retinol is first converted to its ester by the enzyme lecithin:retinol acyltransferase as described above in the RPE, and the products coalesce into lipid droplets, i.e. dynamic organelles termed 'retinosomes'. The next step involves a dual-purpose enzyme (RPE65) in the endoplasmic reticulum, which cleaves the O-alkyl bond (not a conventional hydrolysis reaction) in the retinol ester and at the same time causes a change in the geometry of the double bond in position 11 of retinol from trans to cis. The 11-cis-retinol is then oxidized to 11-cis-retinal by 11-cis-retinal dehydrogenase (RDH5). The full cycle is shown in Figure \(18\ below. The final part of the cycle occurs in the photoreceptor, where first the 11-cis-retinal is reacted with opsin to produce the protein conjugate rhodopsin in a protonated form. When rhodopsin is activated by light, the cis-double bond in the retinoid component is isomerized non-enzymatically by the energy of a photon to the 11‑trans form with a change of conformation that in turn affects the permeability of the membrane and influences calcium transport. This results in further molecular changes that culminate in the release of opsin and all-trans-retinal, which is the trigger that sets off the nerve impulse so that the light is perceived by the brain. A second mechanism for 11-cis-retinal formation that may function to ensure continuous visual responsiveness in bright light involves the (RPE)-retinal G protein-coupled receptor (RGR), which can function as a retinaldehyde photoisomerase. As the enzyme RPE65 functions optimally under low light conditions, it is believed that RGR prevents the saturation of photoreceptors under high light levels, and in this way facilitates vision in daylight. The isomerase, RPE65, and the photoisomerase, RGR, operate together to provide a sustained supply of the visual chromophore under different levels of illumination. The all-trans-retinal is removed from the photoreceptor either by reduction to all-trans-retinol by all-trans-retinol dehydrogenase 8 expressed in the outer segments of photoreceptors or after transport by means of a specific transporter (ABCA4), which provides phosphatidylethanolamine (PE) for conversion to the Schiff-base adduct, i.e. N-retinylidene-phosphatidylethanolamine, as shown in Figure \(19\). It flips from the lumen to the cytosolic leaflet of the disc membrane. This process prevents non-specific aldehyde activity with the effect of removing potentially toxic retinoid compounds from the photoreceptors. The adduct is a transient sink that dissociates so the retinal can be reduced back to all-trans-retinol by the cytoplasmic retinol dehydrogenase. All-trans-retinol exits the photoreceptor and enters the retinal pigment epithelium with the aid of binding to the retinoid-binding protein (IRBP) where it is converted back to a retinyl ester to complete the cycle and restore light sensitivity. As a side-reaction, some troublesome bis-retinoid adducts of PE (and further byproducts) may be produced by non-enzymatic mechanisms, and these can accumulate with age to affect vision. Lower organisms: Bacteriorhodopsin is the best studied of a family of opsins, found in archaea, eubacteria, fungi, and algae. It is a protein with seven transmembrane helices that acts as an opto-electrical transducer or light-gated active ion pump to capture photon energy via its covalently bound chromophore, all-trans-retinal, converting it to 13-cis-retinal, and moves protons against their electrochemical gradient from the cytoplasm to the extracellular space. In Archaea, it is known as the "purple membrane" and can occupy a high proportion of the surface area of the organism. Other Functions of Retinoids in Health and Disease In addition to their function in vision, it is now realized that retinoids have essential roles in growth and development, reproduction and resistance to infection. They are particularly important for the function of epithelial cells in the digestive tract, lungs, nervous system, immune system, skin, and bone at all stages of life. They are required for the regeneration of damaged tissues, including the heart, and they appear to have some potential as chemo-preventive agents for cancer and for the treatment of skin diseases such as acne. Under pathological conditions, stellate cells lose their retinoid content and transform into fibroblast-like cells, contributing to the fibrogenic response. Cirrhosis of the liver is accompanied by a massive loss of retinoids, but it is not clear whether this is a cause or a symptom, and there appears to be confusion as to when supplementation may be helpful in this and other diseases of the liver. Like retinol and retinoic acid, the metabolite 9-cis-retinoic acid also has valuable pharmaceutical properties. With such a large number of double bonds in conjugation, it is not surprising that carotenoids in general, and retinoids in particular are efficient quenchers of singlet oxygen and scavengers of other reactive oxygen species. However, any direct antioxidant properties are not believed to be important in terms of general health in vivo, and it is not clear how relevant the physical properties of retinoids are to specific biochemical processes in comparison to their effects on signaling and gene transcription. There is a caveat that retinoids may stimulate some antioxidant genes and so have an indirect antioxidant function. In fact, nutritional studies with dietary supplements of carotenoids have sometimes suggested pro-oxidant activity. One explanation for detrimental effects may be that regeneration of the parent carotenoid or retinoid from the corresponding radical cation may be limited when concentrations of reductants such as ascorbic acid are low. Many of the retinol metabolites function as ligands to activate specific transcription factors for particular receptors in the nucleus of the cell, and thus they control the expression of a large number of genes (>500), including those essential to the maintenance of normal cell proliferation and differentiation, embryogenesis, for a healthy immune system, and for male and female reproduction. In the innate immune system, vitamin A is required for the differentiation of cells such as macrophages, neutrophils and natural killer cells, while all-trans-retinoic acid is involved in differentiating the precursors of dendritic cells. Retinoic acid and its 9-cis-isomer are especially important in this context, and they are often considered the most important retinoids in terms of function other than in the eye. This structure of the cis-isomer is shown in Figure \(20\). In essence, retinoic acid moves to the nucleus with the aid of small intracellular lipid-binding proteins (CRABP2 and FABP5), which channel it to specific nuclear receptors, the retinoic acid receptors (RAR) of which there are three, RAR-α, β and γ. These are ligand-dependent regulators of transcription and they function in vivo as heterodimers with retinoid X receptors (RXR) to process the retinoic acid signal by acting through polymorphic retinoic acid response elements (RAREs) within the promoter regions of responsive genes. Similarly, 9-cis-retinoic acid and 9-cis-13,14-dihydroretinoic acid are high-affinity ligands for RXR in mice. Together with retinoic acid, these are also ligands for the farnesoid X receptor (FXR), which forms a heterodimer with RXR. The latter receptor complex is involved primarily in bile acid homeostasis, and conversely, there are suggestions that bile acids may have regulatory effects on vitamin A homeostasis. In addition, other nuclear receptors, such as the peroxisome proliferator-activated receptor PPARγ forms a heterodimer with the retinoid X receptor and is activated by retinoic acid to recruit cofactors. This complex in turn binds to the peroxisome proliferator response element (PPRE) gene promoter, leading to regulation mainly of those genes involved in lipid and glucose metabolism, including some involved in inflammation and cancer. To add to the complexity, retinoic acid has extra-nuclear, non-transcriptional effects, such as the activation of protein kinases and other signaling pathways. It has also become evident that many of the functions of retinoids are mediated via the action of specific binding proteins (as discussed briefly above), which control their metabolism in vivo by reducing the effective or free retinoid concentrations, by protecting them from unwanted chemical attack, and by presenting them to enzyme systems in an appropriate conformation. With some tissues, retinol-bound RBP in the blood is recognized by the membrane protein STRA6, which transports retinol into cells where it binds to an intracellular retinol acceptor, cellular retinol-binding protein 1 (CRBP1), and is then able to activate a signaling cascade that targets specific genes. In addition, a specific retinol-binding protein secreted by adipose tissue (RPB4) is involved in the development of insulin resistance and type 2 diabetes, possibly by affecting glucose utilization by muscle tissue, with obvious application to controlling obesity. In the eye, the activity of retinoic acid during development is controlled by binding to apolipoprotein A1. All-trans-retinoic acid has been shown to be effective against many different types of human cancers, especially in model systems but also in some clinical trials, because of its specific effects on cell proliferation, differentiation, and apoptosis (where its relatively low toxicity at normal tissue levels is a virtue). For example, it induces complete remission in most cases of acute promyelocytic leukemia when administered in combination with other chemotherapy techniques. Similarly, 13-cis-retinoic acid has been used successfully in the treatment of children with high-risk neuroblastoma to reduce the risk of recurrence and increase long-term survival rates. However, the efficacy of similar treatments against other types of acute myeloid leukemia and solid tumors appears to be poor. It is hoped that current efforts to obtain a better understanding of the mechanism of the anti-cancer activities will lead to improved treatments. Synthetic analogs of retinoic acid, termed rexinoids, which activate retinoic X receptors, also hold promise as anti-cancer agents. Vitamin A deficiency in children and adult patients is usually accompanied by impairment of the immune system, leading to a greater susceptibility to infection and an increased mortality rate, often with growth retardation and congenital malformations. However, vitamin A deficiency in malnourished children is the major reason for childhood mortality in the underdeveloped world, causing over 650,000 early childhood deaths annually and pediatric blindness. This is doubly tragic in that it is so easily prevented. In adults, vitamin A deprivation affects the reproductive system, inhibiting spermatogenesis in males and ovulation in females. Unfortunately, it is not always easy to distinguish between the effects of vitamin A deficiency and primary defects of retinoid signaling. Functions of Xanthophylls and Other Carotenoids in Humans Xanthophylls are plant C40 tetraterpenes that differ from the carotenoids in having oxygen atoms in the ring structures (hydroxyl, oxo, or epoxyl). Lutein, zeaxanthin, and meso-zeaxanthin from dietary sources, such as green leafy vegetables and yellow and orange fruits and vegetables, are found specifically in the macula of the eye in humans and other primates, i.e. the functional center of the retina in a small central pit known as the macula lutea, where they enhance visual acuity and protect the eye from high-intensity, short-wavelength visible light. They are powerful antioxidants in a region vulnerable to light-induced oxidative stress. Binding proteins specific for lutein- and zeaxanthin mediate the highly selective uptake of these carotenoids into the retina, but meso-zeaxanthin is mainly a metabolite of dietary lutein. Macular xanthophylls decrease the risk of age-related macular degeneration. In the brain, they may stimulate and maintain cognitive function in the elderly, and assist with brain development in infants. Hydroxylated xanthophylls such as lutein, shown in Figure \(21\), occur both in the free form and esterified to fatty acids; the latter are hydrolyzed in the intestines when consumed by animals. Many other carotenoids are absorbed from the diet, and are subject to oxidative cleavage or other catabolic processes, partly in the intestines and partly in other tissues after transport in the lipoproteins. Some carotenoids remain intact and are believed to act as antioxidants, and some may have specific anti-inflammatory actions. For example, carotenoids accumulate in the skin of mammals, where they may have an antioxidant and photo-protective role as well as effects on the moisture content, texture, and elasticity. Lycopene may have protective effects against atherogenesis, coronary heart disease, and prostate cancer. Functions of Carotenoids in Plants As carotenoids have a polyene chain of 9 to 11 double bonds in conjugation, they are able to absorb light in the gap of chlorophyll absorption, and so function as additional light-harvesting pigments in plants. Their distinctive arrangement of electronic levels gives them the capacity to transfer excitation energy from the carotenoid excited state to chlorophyll in the light-harvesting complex (photosystem II). Energy can also be transferred back from chlorophyll to carotenoids as a photoprotection mechanism. During photosynthesis, damaging species are produced by both light and oxygen with reactive oxygen species (ROS) of special concern. The energy is transferred from chlorophyll to the polyene tail of the carotenoid where electrons are moved between the carotenoid bonds until the most balanced or lowest energy state (state) is reached. While there is therefore appreciable potential for carotenoids to act as antioxidants in plants, it is uncertain how important this is from a practical functional standpoint. The length of the polyene tail of carotenoids determines which wavelengths of light will be absorbed by the plant, and those not absorbed are reflected and so determine coloration. F Carotenoids are precursors for two plant hormones and a diverse set of apocarotenoids. For example, abscisic acid, shown in Figure \(21\), is a C15 isoprenoid plant hormone, which is synthesized in plastids from the C40 carotenoid zeaxanthin. A series of enzyme-catalyzed epoxidations and isomerizations is involved followed by cleavage of the intermediate product by a dioxygenation reaction and further oxidations to yield eventually abscisic acid. Functioning via signaling cascades, abscisic acid regulates innumerable biological effects in plants, especially in relation to developmental processes that include plant growth, seed and bud dormancy, embryo maturation and germination, cell division and elongation, floral growth and the control of stomatal closure. It is critical for the responses to environmental stresses that include drought, cold and heat stress, salinity, and tolerance of heavy metal ions. Similarly, strigolactones are C15 oxidation products of carotenoids that are involved in the regulation of symbiosis between plants and arbuscular mycorrhizal fungi and in interactions with plant parasites. Isoprenoids: 3. Other Membrane-Associated Isoprenoids Terpenes (isoprenoids) are one of the most varied and abundant natural products produced by animals, plants, and bacteria. They are generally defined on the basis of their biosynthetic derivation from isoprene units (C5H8), with 55,000 different types characterized to date according to a recent estimate. By most definitions, all isoprenoids should be classified as ‘lipids’, from simple monoterpenes such as geraniol, which is derived from two prenol units, to complex polymers such as natural rubber. Only those isoprenoids that have a functional role in cellular membranes will be discussed here. These include plastoquinone, ubiquinone (coenzyme Q), phylloquinone and menaquinone (vitamin K), dolichol and polyprenols, undecaprenyl phosphate and lipid II, and farnesyl pyrophosphate, together with some key biosynthetic precursors. The nature and function of tocopherols and tocotrienols (vitamin E) and retinoids (vitamin A) are relevant here, but have been discussed previously. Of course, sterols are also isoprenoids. There are two basic mechanisms for the biosynthesis of the isoprene units that are the precursors for the biosynthesis of isoprenoids, i.e., isopentenyl pyrophosphate and dimethylallyl pyrophosphate. These are the mevalonate pathway, which is located in the cytosol of the cell, and the non-mevalonate pathway, found mainly in the plastids of plants. These have been discussed previously. Phytol Phytol or (2E,7R,11R)-3,7,11,15-tetramethyl-2-hexadecen-1-ol, i.e., with 20 carbons in a 16-carbon chain and one double bond, is an acyclic diterpene alcohol, which is synthesized in large amounts in plants as an essential component of chlorophyll, the most important photosynthetic pigment in plants and algae. Geranylgeranyl-diphosphate synthesized in chloroplasts via the 4-methylerythritol-5-phosphate (non-mevalonate) pathway is the primary precursor of phytol following reduction of three double bonds by geranylgeranyl reductase, and this can occur before or after attachment to chlorophyll, depending upon species. Chlorophyll dephytylase (CLD1) is the enzyme in plants responsible for chlorophyll hydrolysis and the release of phytol. The reactions are shown in Figure \(22\). Little free phytol is present in plant tissues, although some phytol esters of fatty acids may occur, especially when plants are stressed during nitrogen deprivation or in senescence, when chlorophyll is degraded, fatty acids are released from glycerolipids and a phytol ester synthase is induced as part of a detoxification and recycling process. Bell peppers and rocket salad are especially rich sources under normal conditions. In addition, phytol as its diphosphate is utilized for the synthesis of tocopherols (vitamin E) and phylloquinol (vitamin K - see below), and the precursor geranylgeraniol and its fatty acid ester occur in small amounts in some plant species. It is the biosynthetic precursor of tocotrienols and the highly unsaturated carotenoids (and hence of retinoids). Phytenal has been isolated as an intermediate in the catabolism of phytol in plants, but further steps are uncertain although phytanoyl-CoA has been detected in stressed plants. As phytenal is highly reactive and potentially toxic via its interaction with proteins, its accumulation must be kept at a low level by competing pathways. In ruminant animals, chlorophyll is hydrolyzed by rumen microorganisms with the release of free phytol. This does not occur in humans, but some phytol may be ingested with plant foods either in free form or as phytol esters and can be absorbed from the intestines. Within animal tissues, phytol is oxidized to phytanic acid. Phytol and/or its metabolites have been reported to activate the transcription factors PPARα and retinoid X receptor. In mice, oral phytol induces a substantial proliferation of peroxisomes in many organs. Plastoquinone A molecule that is related to the tocopherols, plastoquinone, is found in cyanobacteria and plant chloroplasts, and it is produced in plants by analogous biosynthetic pathways to those of tocopherols in the inner chloroplast envelope with solanesol diphosphate as the biosynthetic precursor of the side chain; there appears to be a somewhat different mechanism in cyanobacteria. The molecule is sometimes designated - 'plastoquinone-n' (or PQ-n), where 'n' is the number of isoprene units, which can vary from 6 to 9. It's structure is shown in Figure \(23\). Plastoquinone has a key role in photosynthesis, by providing an electronic connection between photosystems I and II, generating an electrochemical proton gradient across the thylakoid membrane. This provides energy for the synthesis of adenosine triphosphate (ATP). The reduced dihydroplastoquinone (plastoquinol) that results in the transfers further electrons to the photosynthesis enzymes before being re-oxidized by a specific cytochrome complex; the redox state of the plastoquinone pool regulates the expression of many of the genes encoding photosystem proteins. X-Ray crystallography studies of photosystem II from cyanobacteria show two molecules of plastoquinone forming two membrane-spanning branches. In addition, plastoquinone has antioxidant activity comparable to that of the tocopherols, protecting especially against excess light energy and photooxidative damage. Similarly, in thylakoid membranes, plastoquinol is able to scavenge superoxide with the production of H2O2. Plastoquinone is a cofactor participating in desaturation of phytoene in carotenoid biosynthesis, and the biosynthetic precursor of plastochromanols. With these many different functions, plastoquinone connects photosynthesis in plants with metabolism, light acclimation, and stress tolerance. Plastoquinone-9, together with phylloquinone, tocopherol, and plastochromanol-8, is stored in plastoglobuli, lipoprotein-like micro-compartments, which enable exchange with the thylakoid membrane and are also involved in chlorophyll catabolism and recycling. It has been suggested that the redox state of the plastoquinone pool is the main redox sensor in chloroplasts that initiates many physiological responses to changes in the environment and in particular to those related to light intensity by regulating the expression of chloroplast genes. Ubiquinone (Coenzyme Q) The ubiquinones, which are also known as coenzyme Q (CoQ) or mitoquinones, have obvious biosynthetic and functional relationships to plastoquinone and they are found in all the domains of life (hence the name). They have a 2,3‑dimethoxy-5-methylbenzoquinone nucleus and a side chain of six to ten isoprenoid units; the human form illustrated below has ten such units (coenzyme Q10), i.e., it is 2,3-dimethoxy-5-methyl-6-decaprenyl-1,4-benzoquinone, while that of the rat has nine, Escherichia coli has eight and Saccharomyces cerevisiae has six. In plants, ubiquinones tend to have nine or ten isoprenoid units. Forms with a second chromanol ring, resembling the structures of tocopherols, are also produced (ubichromanols), but not in animal tissues. They are generated on an industrial scale for pharmaceutical purposes by yeast fermentation. Because of their hydrophobic properties, ubiquinones are located entirely in membrane bilayers in most eukaryote organelles, probably at the mid-plane. Ubiquinones are synthesized de novo in mitochondria in most cells in animal, plant, and bacterial tissues by a complex sequence of reactions from the essential amino acid phenylalanine and then tyrosine to generate p-hydroxybenzoic acid, which is the key precursor that is condensed with the polyprenyl unit (from the cholesterol synthesis pathway) via a specific transferase; this is followed by decarboxylation, hydroxylation, and methylation steps, depending on the specific organism, although some of the required enzymes have yet to be fully characterized. In Escherichia coli, biosynthesis does not occur in a membrane environment as had been thought. Rather, the seven proteins that catalyze the last six reactions of the biosynthetic pathway, following the attachment of the isoprenoid tail, form a stable complex or metabolon in the cytoplasm so enabling modification of the hydrophobic substrates in a hydrophilic environment. In mitochondria, coenzyme Q is present both as the oxidized (ubiquinone) and reduced (ubiquinol) forms as shown in Figure \(24\). Figure \(24\): Ubiquinone-ubiquinol interconversion Ubiquinones are essential components of the respiratory electron transport system, possibly as part of supramolecular complexes, taking part in the oxidation of succinate or NADH via the cytochrome system to generate the protonmotive force used by the mitochondrial ATPase to synthesize ATP. In this process, coenzyme Q transfers electrons from the various primary donors, including complex I, complex II, and the oxidation of fatty acids and branched-chain amino acids, to the oxidase system (complex III), while simultaneously transferring protons to the outside of the mitochondrial membrane with the result of a proton gradient across the membrane. As a consequence, it is reduced to ubiquinol. Thus, it is an essential component of the cycle that generates the proton motive force driving ATP production via oxidative phosphorylation. In yeast, one coenzyme Q binding protein (COQ10), and in humans two related proteins (COQ10A and COQ10B) may serve as chaperones or transporters during this process. Mitochondrial coenzyme Q is also implicated in the production of reactive oxygen species by a mechanism involving the formation of superoxide from ubisemiquinone radicals, and in this way is responsible for causing some of the oxidative damage behind many degenerative diseases. In this action, it is a pro-oxidant. It is most abundant in organs with a high metabolic rate such as the heart, kidneys, and liver. In complete contrast in its reduced form (ubiquinol) in non-mitochondrial cellular membranes and plasma lipoproteins, it acts as an endogenous antioxidant, the only lipid-soluble antioxidant to be synthesized endogenously. It inhibits lipid peroxidation in biological membranes and serum low-density lipoproteins, and it may also protect membrane proteins and DNA against oxidative damage. The ferroptosis suppressor protein 1 (FSP1) replenishes ubiquinol, and this acts protectively by combating the lipid peroxidation that drives ferroptosis. The mechanism involves the recruitment of FSP1 to the plasma membrane following myristoylation, where this functions as an oxidoreductase that reduces ubiquinone to ubiquinol, which acts as a lipophilic radical-trapping antioxidant that halts the propagation of lipid peroxides. In this manner, it regulates cellular redox status and cytosolic oxidative stress, and thereby is a controlling factor in apoptosis. Other NAD(P)H dehydrogenases with CoQ reductase activity include cytochrome b5 reductase and NQo1 (NAD(P)H:quinone oxidoreductase). Although ubiquinone has only about one-tenth of the antioxidant activity of vitamin E (α-tocopherol), it is able to stimulate the effects of the latter by regenerating it from its oxidized form back to its active fully reduced state (similarly with vitamin C). However, ubiquinones and tocopherols appear to exhibit both cooperative and competitive effects under different conditions. Similarly, in bacteria and other prokaryotes, ubiquinones participate a large number of redox reactions, notably in the respiratory electron transport system but also in other enzyme reactions that require electron donation, including the formation of disulfide bonds. Coenzyme Q has many other functions that are not related directly to its antioxidant function. Some coenzyme Q is used in mitochondria by enzymes that link the mitochondrial respiratory chain to other metabolic pathways, including fatty acid β-oxidation as an electron acceptor, nucleotide biosynthesis de novo, amino acid oxidation (glycine, proline, glyoxylate, and arginine), and detoxification of sulfide. Via these activities, coenzyme Q may modulate metabolic pathways located outside the mitochondria indirectly. There are also suggestions that coenzyme Q may be involved in redox control of cell signaling and gene expression, and in particular to repress the expression of inflammatory genes. In relation to its anti-inflammatory properties, clinical studies suggest that supplementation with coenzyme Q10 reduces the levels of the inflammatory mediators C-reactive protein, interleukin-6, and tumor necrosis factor alpha (TNFα) to a significant degree. In addition, it is a regulator of mitochondrial permeability, and in relation to pyrimidine nucleotide biosynthesis, it is required for DNA replication and repair. Dietary ubiquinone, i.e., that in food or dietary supplements, is absorbed by enterocytes via a process of “passive facilitated diffusion”, probably requiring a carrier molecule, before incorporation into chylomicrons for transport to the liver. This eventually leads to elevated levels of ubiquinol in blood, especially in the LDL and VLDL lipoproteins, presumably because of the reduction of the oxidized form in the lymphatic system. In consequence, there is reported to be enhanced protection against lipid peroxidation with beneficial effects on health, especially in relation to cardiac function, sperm motility, and neurodegenerative diseases. For example, CoQ levels in both plasma and the heart correlate with heart failure in patients, and clinical trials of dietary supplementation have shown promising results. This may be of particular importance in the elderly or in patients on statins, when endogenous synthesis declines. A CoQ10 deficiency syndrome is associated with inherited pathological diseases, defined by a decrease of the CoQ10 content in muscle and/or cultured skin fibroblasts. Early clinical trials with CoQ10 and a synthetic analog, idebenone, against various neurodegenerative diseases, including Alzheimer's disease, Parkinson's disease, Huntington's disease, and others, are encouraging. Phylloquinone and Menaquinones (Vitamin K) Phylloquinone or 2-methyl-3-phytyl-1,4-naphthoquinone is synthesized in the inner chloroplast envelope of cyanobacteria, algae, and higher plants by a mechanism analogous to that of the tocopherols, i.e., from chorismate in the shikimate pathway with a prenyl side chain derived from phytyldiphosphate. In this membrane, it is a key component of the photosystem I complex where it receives an electron from the chlorophyll a acceptor molecule and then donates an electron to the membrane-associated iron-sulfur protein acceptor cluster in the complex. In an obvious parallel to the plastoquinones (above), two molecules of phylloquinone form two membrane-spanning branches, as demonstrated by X-ray crystallography studies of photosystem I from cyanobacteria. Plastoglobules associated with the thylakoid membrane are believed to function as a reservoir for excess phylloquinone, and may also function in its metabolism. Edward A. Doisy and Henrik Dam received the Nobel Prize in Physiology or Medicine in 1943 for their discovery of vitamin K and its chemical structure. The structure of vitamin K is shown in Figure \(25\). The menaquinones are related bacterial products, which function in the respiratory and photosynthetic electron transport chains of bacteria. They have a variable number (4 to 10) of isoprenoid units in the tail, and they are sometimes designated ‘MK-4’ to ‘MK-10’. In contrast to phylloquinone, these are usually highly unsaturated. In some species, there are methyl or other groups attached to the naphthoquinone moiety. Remarkably high concentrations of menaquinones are present in membranes of some extremophiles such as the haloarchaea, where it has been suggested that they act as ion permeability barriers and as a powerful shield against oxidative stress in addition to their functions as electron and proton transporters. Phylloquinone is an essential component of the diet of animals and has been termed 'vitamin K1'. It must be supplied by green plant tissues, where it occurs in the range 400-700 μg/100 g, or seed oils. The menaquinones, the main source of which in the human diet is cheese and yogurt, also have vitamin K activity and are termed 'vitamin K2'. They account for about 10-25% of the vitamin K content of the Western diet. A synthetic saturated form of this, which is used in animal feeds, is known as 'vitamin K3 or menadione', though strictly speaking it is not a vitamin but a pro-vitamin in that it can be converted to the menaquinone MK-4 in animal tissues by addition of a phytyl unit; it is too toxic for human nutrition. Vitamin K forms are absorbed from the intestines and transported in plasma in the form of lipoproteins in a similar manner to the other fat-soluble vitamins. Different tissues have differ storage capacities and presumably requirements for the various forms of vitamin K. As a high proportion is excreted, there appears to be a requirement for a constant intake. Vitamin K1 is taken up rapidly by the liver, but vitamin K2 remains in the plasma for much longer and maybe the main source of the vitamin in peripheral tissues. In addition, it has been established that some dietary phylloquinone is converted to menaquinone-4 in animals, although the quantitative significance has still to be established; the mechanism involves conversion to menadione in the intestines followed by transport to tissues where a geranylgeranyl side-chain is attached by a specific prenyl transferase. The primary role of vitamin K in animal tissues is to act as a cofactor specific to the vitamin K-dependent enzyme γ-glutamyl carboxylase in the endoplasmic reticulum in the liver mainly. Its function is the post-translational carboxylation of glutamate residues to form γ-carboxyglutamic acid in proteins, such as prothrombin. In this way, prothrombin and three related proteins are activated to promote blood clotting. The γ-carboxyglutamic acid residues are located at the binding site for Ca2+, and are vital for the activity of the enzyme. Phylloquinone must first be converted to the reduced form, phylloquinol, which is the actual cofactor for the enzyme; molecular oxygen and carbon dioxide are both required also. Phylloquinol donates hydrogen to the glutamic acid residue and is oxidized in the process to 2,3-epoxyphylloquinone. A further enzyme, vitamin K epoxide reductase, regenerates phylloquinone by reduction of the epoxide in a dithiol-dependent reaction so that this can be re-utilized many times ("the vitamin K cycle", shown below in Figure \(26\)). Menaquinones undergo the same cycle of reaction. By interfering with the last step in the metabolic cycle, warfarin, the rodenticide, prevents blood clotting. In the same way, a deficiency in vitamin K results in the inhibition of blood clotting and can lead to brain hemorrhaging in malnourished newborn infants, though this is not seen in adult humans, presumably because intestinal bacteria produce sufficient for our needs. Vitamin K-dependent proteins are also known to have important functions in the central and peripheral nervous systems, and vitamin K influences sphingolipid biosynthesis in the brain. Unsaturated isoprene units rather than phytol are used for the biosynthesis of menaquinones, and they differ from phylloquinone with respect to their chemical structure and pharmacokinetics. It is now apparent that vitamin K2 (MK-4 especially) has a number of different actions, some with specificities for particular tissues. For example, osteocalcin is a γ-carboxyglutamic acid-containing protein, which forms a strong complex with the mineral hydroxyapatite (calcium phosphate) of bone; it must be carboxylated to function properly and vitamin K2 appears to be of particular importance in this instance. Vitamin K2 also regulates bone remodeling by osteoclasts to remove old or damaged bone and its replacement by new bone. In addition, vitamin K2 is involved in vascular calcification, cell growth, and apoptosis. Side effects of the use of anticoagulants that bind to vitamin K can be osteoporosis and increased risk of vascular calcification. Although careful control of the dosage is necessary, vitamin K2 may be a useful adjunct for the treatment of osteoporosis, and it may reduce morbidity and mortality in cardiovascular health by reducing vascular calcification. Excess vitamin K1 and the menaquinones are catabolized in the liver by a common degradative pathway in which the isoprenoid side chain is shortened to yield carboxylic acid aglycones such as menadiol, which can be excreted in bile and urine as glucuronides or sulfates. Dolichols and Polyprenols Polyisoprenoid alcohols, such as dolichols, are ubiquitous if minor components, relative to the glycerolipids, of membranes of most living organisms from bacteria to mammals. They are hydrophobic linear polymers, consisting of up to twenty isoprene residues or a hundred carbon atoms (or many more in plants especially), linked head-to-tail, with a hydroxy group at one end (α-residue) and a hydrogen atom at the other (ω-end). In dolichols (or dihydropolyprenols), the double bond in the α-residue is hydrogenated, and this distinguishes them from the polyprenols with a double bond in the α-residue. Their structures are shown in Figure \(27\). Polyisoprenoid alcohols are further differentiated by the geometrical configuration of the double bonds into three subgroups, i.e., di-trans-poly-cis, tri-trans-poly-cis, and all-trans. For many years, it was assumed that polyprenols were only present in bacteria and plants, especially photosynthetic tissues, while dolichols were found in mammals or yeasts, but it is now known that dolichols can also occur at low levels in bacteria and plants, while polyprenols have been detected in animal cells. Solanesol is a related and distinctive plant product with trans double bonds only and is a precursor of plastoquinone. Within a given species, components of one chain length may predominate, but other homologs are usually present. The chain length of the main polyisoprenoid alcohols varies from 11 isoprene units in eubacteria, to 16 or 17 in Drosophila, 15 and 16 in yeasts, 19 in hamsters, and 20 in pigs and humans. In plants, the range is from 8 to 22 units, but some species of plant have an additional class of polyprenols with up to 40 units. In tissues, polyisoprenoid alcohols can be present in the free form, esterified with acetate or fatty acids, phosphorylated or monoglycosylated phosphorylated (various forms), depending on species and tissue. Polyisoprenoid alcohols per se do not form bilayers in aqueous solution, but rather a type of lamellar structure. However, they are found in most membranes, especially the plasma membrane of liver cells and the chloroplasts of plants. Dolichoic acids, i.e., related molecules with a terminal carboxyl group and containing 14–20 isoprene units, have been isolated from the substantia nigra of the human brain. However, they were barely detectable in the pig brain. Biosynthesis of the basic building block of dolichols, e.g. isopentenyl diphosphate, follows either the mevalonate pathway or a more recently described methylerythritol phosphate pathway discussed in relation to cholesterol biosynthesis previously. Farnesyl pyrophosphate is the primary precursor in the biosynthesis of polyprenols and is the branch point in sterol/isoprene biosynthesis, depending on the nature of the organism (see a further note below). Subsequent formation of the linear prenyl chain is accomplished by cis-prenyl transferases that catalyze the condensation of isopentenyl pyrophosphate and farnesyl pyrophosphate and then the growing the allylic prenyl diphosphate chain. The end products are polyprenyl pyrophosphates, which are dephosphorylated first to polyprenol phosphate and thence to the free alcohol. Finally, a specific reductase has been identified from human tissues that catalyzes the reduction of the double bond in position 2 to produce dolichols. There is a family of cis-prenyl transferases, present in both eukaryotes and bacteria, that in addition to the synthesis of dolichols can catalyze the formation of isoprenoid carbon skeletons from neryl pyrophosphate (C10) to natural rubber (C>10,000), including the polyisoprenoid phosphates involved in protein glycosylation as discussed in the next section. The reactions in the synthesis of dolichol are shown in Figure \(28\). Although polyprenols and dolichols were first considered to be simply secondary metabolites, they are now known to have important biological functions. Glycosylation of asparagine residues is the main protein modification in all three domains of life, and phosphorylated polyisoprenoids, including dolichols, are essential to this process (next section). There is also a suggestion that free dolichol may have a beneficial antioxidant function in cell membranes. Polyisoprenoid Phosphates and Glycosylation of Proteins Glycosylated phosphopolyisoprenoid alcohols are the carriers of oligosaccharide units for transfer to proteins and as glycosyl donors, i.e., they are substrates for glycosyl transferases for the biosynthesis of glycans in a similar manner to the cytosolic sugar nucleotides. They differ from the latter in their intracellular location, with the lipid portion in the membrane of the endoplasmic reticulum and the oligosaccharide portion specifically located either on the cytosolic or lumenal face of the membrane. The degree of unsaturation and chain length of the product is important for recognition by the enzymes in the next stage of the pathway. Dolichol phosphates: In eukaryotes, N-glycosylation begins on the cytoplasmic side of the endoplasmic reticulum with the transfer of carbohydrate moieties from nucleotide-activated sugar donors, such as uridine diphosphate N-acetylglucosamine, onto dolichol phosphate. Then, N-acetylglucosamine phosphate is added to give dolichol-pyrophosphate linked to N-acetylglucosamine, to which a further N-acetylglucosamine unit is added followed by five mannose units, the last catalyzed by dolichol phosphate mannose synthase, which is also essential for GPI-anchor biosynthesis. The dolichol-PP glycan is shown in Figure \(29\). The resulting dolichol-pyrophosphate heptasaccharide is then flipped across the endoplasmic reticulum membrane to the luminal face with the aid of a “flippase”. Four further mannose and three glucose residues are added to the oligosaccharide chain by means of glycosyltransferases, which utilize as donors dolichol-phospho-mannose and dolichol-phospho-glucose, which are also synthesized on the cytosolic face of the membrane and flipped across to the luminal face. In humans, the final lipid product is a C95-dolichol pyrophosphate-linked tetradecasaccharide, the oligosaccharide unit of which is transferred from the dolichol carrier onto specific asparagine residues on a developing polypeptide in the membrane. The carrier dolichol-pyrophosphate is dephosphorylated to dolichol-phosphate then diffuses or is flipped back across the endoplasmic reticulum to the cytoplasmic face. The Archaea use dolichol in their synthesis of lipid-linked oligosaccharide donors with both dolichol phosphate (Euryarchaeota) and pyrophosphate (Crenarchaeota) as carriers; these can have variable numbers of isoprene units many of which can be saturated. In the haloarchaeon Haloferax volcanii, for example, a series of C55 and C60 dolichol phosphates with saturated isoprene subunits at the α- and ω-positions is involved in the glycosylation reaction of target proteins, while similar lipid carriers of oligosaccharide units appear to be present in methanogens. Archaea of course use isoprenyl ethers linked to glycerol as major membrane lipid components in addition to unusual carotenoids such as the C50 bacterioruberins. In many of these species, isoprenoid biosynthesis is via the 'classical' mevalonate pathway, but in other species, some aspects of this pathway differ. Undecaprenylphosphate: Most other bacteria use undecaprenyl-diphosphate-oligosaccharide as a glycosylation agent in a similar way for the biosynthesis of peptidoglycan, the main component of most bacterial cell walls and a structure unique to bacteria, of many other cell-wall polysaccharides, including lipopolysaccharides, O-antigenic polysaccharides and capsular polysaccharides, and of N-linked protein glycosylation in both in Gram-negative and Gram-positive bacteria. Undecaprenyl phosphate (a C55 isoprenoid), also referred to as bactoprenol, is the essential lipid intermediate. Its structure is shown in Figure \(30\). It differs from the dolichol phosphates mainly in that the terminal unit is unsaturated, and is synthesized by the addition of eight units of isopentenyl pyrophosphate to farnesyl pyrophosphate, a reaction catalyzed by undecaprenyl pyrophosphate synthase in the cytoplasm, followed by the removal of a phosphate group. Undecaprenyl phosphate is required for the synthesis and transport of glycans for external polymer formation. Thus, glycans are covalently transferred to the carrier lipid by membrane-embedded or membrane-associated enzymes using nucleotide-activated precursors. For example, the carrier lipid with GlcNAc-MurNAc-peptide monomers, i.e., as lipid II, is hydrophilic and is then transported across the cytoplasmic membrane to external sites for peptidoglycan formation. Similarly, it is required for the synthesis of lipoteichoic acids and lipopolysaccharide O-antigens. Lipid II: Undecaprenyl diphosphate-MurNAc-pentapeptide-GlcNAc, often simply termed lipid II, is the last significant lipid intermediate in the construction of the peptidoglycan cell wall in bacteria (Lipid I is the biosynthetic precursor lacking the N-acetylgluosamine residue). Its structure is shown below in Figure \(31\). Figure \(31\): Lipid II This molecule must be translocated from the cytosolic to the exterior membrane of the organism, and three different protein classes have been identified that can accomplish this of which ATP-binding cassette (ABC) transporters are best characterized. Once across the membrane, lipid II is cleaved to provide the MurNAc-pentapeptide-GlcNAc monomer, which undergoes polymerization and crosslinking to form the complex peptidoglycan polymer that provides strength and shape to bacteria. The undecaprenyl-pyrophosphate remaining is hydrolyzed to undecaprenyl phosphate by a membrane-integrated member of the type II phosphatidic acid phosphatase family and is recycled back to the interior of the membrane by an as yet unidentified transport mechanism. The turnover rate is very high so the lipid II cycle is considered to be the rate-limiting step in peptidoglycan biosynthesis. There is also some evidence that lipid II has a function on the inner leaflet of the cytoplasmic membrane in organizing the proteins of the cytoskeleton. Because of its highly conserved structure and accessibility on the surface membrane, the proteins involved in the synthesis and transport of lipid II are considered important targets for the development of novel antibiotics. Figure \(\PageIndex{xx}\): In a few prokaryotes, the membrane intermediate has a polyprenyl-monophosphate-glycan structure instead of lipid II, and undecaprenyl-phosphate-L-4-amino-4-deoxyarabinose is involved in lipid A modification in Gram-negative bacteria, for example. There are obvious parallels with the involvement of glycosylated phosphopolyisoprenoid alcohols as carriers of oligosaccharide units for transfer to proteins and as glycosyl donors in higher organisms (see above). β‑D‑Mannosyl phosphomycoketide is an isoprenoid phosphoglycolipid found in the cell walls of Mycobacterium tuberculosis, the lipid component of which is a C32-mycoketide, consisting of a saturated oligoisoprenoid chain with five chiral methyl branches. It acts as a potent antigen to activate T-cells upon presentation by CD1c protein. Its structure is shown in Figure \(32\). Farnesyl Pyrophosphate and Related Compounds Farnesyl pyrophosphate is a key intermediate in the biosynthesis of sterols such as cholesterol, and it is the donor of the farnesyl group in the biosynthesis of dolichols and polyprenols (see above) as well as for the isoprenylation of many proteins (see the web page on proteolipids). However, it is also known to mediate various biological reactions in its own right via interaction with a specific receptor. It is synthesized by two successive phosphorylation reactions of farnesol. Presqualene diphosphate is unique among the isoprenoid phosphates in that it contains a cyclopropylcarbinyl ring. In addition to being a biosynthetic precursor of squalene, and thence of cholesterol, it is a natural anti-inflammatory agent, which functions by inhibiting the activity of phospholipase D and the generation of superoxide anions in neutrophils. Their structures are shown in Figure \(32\).

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/21%3A_Lipid_Biosynthesis/21.06%3A_Biosynthesis_of_Isoprenoids.txt

Search Fundamentals of Biochemistry Introduction Organic chemistry is usually described as the chemistry of carbon-containing molecules. But isn't that definition a bit carbon centric, especially since the prevalence of oxygen-containing molecules is staggering? What about nitrogen? We live in a dinitrogen-rich atmosphere (80%), and all classes of biomolecules (lipids, carbohydrates, nucleic acids, and proteins) contain nitrogen. Dinitrogen is very stable, given its triple bond and nonpolarity. We rely on a few organisms to fix N2 from the atmosphere to form ammonium (NH4+), which through nitrification and denitrification can form nitrite (NO2-), nitrate (NO2-), nitric oxide (NO), and nitrous oxide (N2O), the latter being a potent greenhouse gas. We'll concentrate on the metabolic fate of amino groups in amino acids and proteins in the next section. Before exploring their fates, look at Figure $1$ which shows an overall view of the biological nitrogen cycle. The study of biochemistry should encompass more than homo sapiens and expand to the ecosystem in which we are such a small but damaging part. Figure $1$: Nitrogen Cycle Let's break down the diagram from a biochemical perspective. There are aerobic and anaerobic processes (conducted by bacteria). Nitrogen-containing substances include both inorganic (ammonium, nitrate, nitrite) and organic (amino acids, nucleotides, etc) molecules. The reactions shown are oxidative and reductive (note: the oxidation number of the nitrogen atoms in the molecules is shown in red). Most of the reactions are carried out underground by bacterial and Archaeal microorganisms. Here are some of the major reactions: • N2 fixation (a reduction): N2 from the air is converted by bacteria to ammonium (NH4+) by the enzyme nitrogenase of soil prokaryotes. The energetically disfavored reaction requires lots of ATPs. Ammonium once made can then be taken up by primary producers like plants and incorporated into biomolecules such as amino acids, which animals consume. For those who may still believe that people have marginal effects on our biosphere, consider this. We may soon fix more N2 to NH3 through the industrial Born-Haber reaction (used for fertilizer and explosive productions) that is all made by the biosphere. Much of the nitrogen in use comes from the Born-Haber reaction. The excess NH4+ (upwards of 50%) produced industrially and which enters the soil in fertilizers (mostly as NH4NO3) has overwhelmed nature's ability to balance the nitrogen cycle and is not taken up by plants. It is metabolized by microorganisms to nitrite and nitrate. • Nitrification: Ammonium is converted to nitrite by ammonia-oxidizing aerobic microorganisms and further to nitrate by a separate group of nitrite-oxidizing aerobic bacteria. Here are the reactions (Rx 1 and 2) to produce nitrate through a hydroxylamine intermediate, followed by the formation of nitrate (Rx 3). NH3 + O2 + 2e- → NH2OH + H2O Rx 1 NH2OH + H2O → NO2- + 5H+ + 4e- Rx 2 NO2- + 1/2 O2 → NO3- Rx 3 These added ions exceed soil capacity and end up runoff water, polluting our rivers and lakes. • Denitrification: This anaerobic reaction pathway reproduces N2 from nitrate Here is the net reaction: 2NO3- + 10e- + 12H+ → 2N2 + 6H2O • Anammox reaction: This more recently discovered bacterial anaerobic reaction pathway converts ammonium and nitrate to N2. Here is the net reaction NO2- + NH3+ → N2 + 2H2O • Ammonification (not to be confused with mummification) occurs when plants and animals decompose, which returns ammonium to the soil for reuse by plants and microbes. These reactions are shown in the abbreviated Nitrogen Cycle shown in Figure $2$. Figure $2$: Abbreviated Nitrogen Cycle Nitrogen metabolites are nutrients for plants and perhaps the most important nutrients in the regulation of plant growth (primary productivity) and in regulating life diversity in the biosphere. All living organisms require feedstocks to produce energy and as substrates for biosynthetic reactions. Which is used depends on the organism. Plants are primary producers so they use their synthesized carbohydrates for both energy production and biosynthesis. For carnivores, proteins and their derived amino acids are the source of energy (through oxidation) and serve as biosynthetic precursors. For omnivorous organisms, the source of energy depends on the "fed" state. With abundant food resources, carbohydrates, and lipids are the source of energy. Unlike carbohydrates and lipids, which can be stored as glycogen and triacylglycerols for future use, excess protein, and their associated amino acids can not be stored, so amino acids can be eliminated or used for oxidative energy. In the fed state, carbohydrates are the main source, while in the unfed state, lipids take a predominant role. Under starving conditions, the organism's own proteins are broken down and used for oxidative energy production and for any biosynthesis that remains. In diseased states like diabetes, which can be likened to a starving state in the presence of abundant carbohydrates, both lipids and amino acids become the sources of energy. How are amino acids in animals oxidatively metabolized? Many pathways could be used to do so but it would seem logical that NH4+ would be removed and the carbons in the remaining molecule would eventually enter glycolysis or the TCA cycle in the form of ketoacids. NH4+ is toxic in high concentrations. Ammonium is not oxidized to nitrite or nitrates in humans as occurs in the soil by microorganisms. It can be recycled back into nucleotides or amino acids, and excess amounts are eliminated from the organism. Both processes must be highly controlled. We will turn out attention to the oxidation of amino acids in the next section. Nitrogenase: An Introduction Beauty is in the eye of the beholder. As the domain of biochemistry covers the entire biological world, the extent of coverage of a given topic in textbooks can depend, in part, on the interest and experiences of the author(s) presenting the material. Is relevance a metric that should determine coverage? If so, books focused on human or medical biochemistry would surely omit photosynthesis. If topics are selected based on their importance for life, then photosynthesis must surely be covered. If so, then nitrogenase must also be included. If the degree of chemical difficulty for a chemical reaction and the amazing eloquence of the evolved biochemistry solution is considered, then both photosynthesis and nitrogen fixation must be presented. Even though nitrogen fixation is a reductive reaction, it shares strong similarities with the oxygen-evolving complex of photosynthesis. They catalyze enormously important redox reactions that involve an abundant atmospheric gas using a very complicated and unique inorganic metallic cofactor that evolution has selected as uniquely suited for the job. Every first-year student of chemistry can draw the Lewis structure of dinitrogen, N2, which contains a triple bond and a lone pair on each nitrogen. If Lewis structures speak to them, they should be able to state that the triple bond makes N2 extraordinarily stable, thus explaining why we can breathe an atmosphere containing 80% N2 and not die. If they have taken biology, they are also aware that very few biological organisms can utilize N2 as a substrate, as this requires breaking bonds between the nitrogen atoms, a chemical process reserved for nitrogen “fixing” bacteria found in rhizomes of certain plants. Lastly, they probably memorized that high pressure and temperature are needed in the Haber-Bosch process used to convert N2 and H2 to ammonia, NH3. As with any scientific advance, the Haber-Bosch process has brought both harm (it's used for explosive weapons) and good (fertilizers). This process now fixes enough N2 in the form of fertilizers to support half of the world’s population, with nitrogenase supporting the rest. Efforts are underway to genetically modify plants to make nitrogenase, eliminating the need for fertilizers but perhaps creating unforeseen problems of its own. You might be surprised to find out that at room temperature the equilibrium constant favors ammonia formation, hence ΔG0 < 0. The reaction is favored enthalpically as it is exothermic at room temperature. It is disfavored entropically as should be evident from the balanced equation: $\ce{N2(g) + 3H2(g) → 2 NH3(g)}. \nonumber$ The thermodynamic parameters for the reaction (per mol) are ΔH° = –46.2 kJ, ΔS° = –389 J K–1, and ΔG° = –16.4 kJ at 298 K The entropy is negative since the reaction proceeds from 4 molecules to two molecules. From an enthalpy perspective, if you raise the temperature of an exothermic reaction, you drive it in a reverse direction. If you increase the pressure, you shift the equilibrium to the side that has the fewest number of molecules. If the reaction is favored thermodynamically at room temperature, why doesn’t it proceed? This story sounds familiar as this same descriptor applies to the oxidation of organic molecules with dioxygen. There we showed using MO theory that the reaction is kinetically slow. Same with NH3 formation. A superficial way to see this is that we must break bonds in the stable N2 to start the reaction, leading to a high activation energy, and making the reaction kinetics sluggish. One could jump-start the reaction by raising the temperature, but that would slow an exothermic reaction. The Keq (or KD) and ΔG0 are functions of temperature and for this reaction, the reaction becomes disfavored at higher temperatures. The solution Haber found was high pressure, forcing the reaction to the side that has fewer molecules of gas, and high temperature to overcome the activation energy barrier and make the reaction kinetically feasible. A complex metal catalyst (magnetite - Fe3O4 -with metal oxides like CaO and Al2O3 which prevent reduction of the Fe with H2) provides an absorptive surface to bring reagents together and facilitate bond breaking in H2 and N2. In photosynthesis, the oxygen-evolving complex (OEC) with Mn, Fe, S, and Ca is used to oxidize another very stable and ubiquitous molecule, H2O. Now we explore the amazing mechanisms behind the nitrogenase complex which fixes N2 to form NH3 in a reductive fashion. What might be needed to drive this reaction biologically? You might surmise the list to include: • a source of energy, most likely ATP, to facilitate this complex reaction; • a source of electrons as the N atoms move from an oxidation state of 0 in elemental N to 3- in NH3; this source turns out to be a protein called flavodoxin or ferredoxin. Of course, these electrons also have interesting sources before they were in the electron carriers of these proteins; • some pretty amazing metal centers to accept and donate electrons in a controlled way; these centers are mostly FeS clusters with an additional cluster containing molybdenum (Mo). The clusters are named F, P, and M • a source of hydrogen; you might have guessed correctly that it’s not H2 gas (from where would that come?), but H+ ions which are pretty ubiquitously available. • a net reaction that is different that the Haber-Bausch process (N2 + 3H2 → 2 NH3). Here is the actual reaction catalyzed by nitrogenase: $\ce{N2 + 8e^{-} + 16ATP + 8H^{+} → 2NH3 + H2 + 16ADP + 16P_i}. \nonumber$ Let’s think a bit about the reaction. As electrons are added the attraction between the nitrogen atoms must decrease. Eventually, bonds between them must be broken. Protons could be easily added to maintain charge neutrality. A basic mechanism might involve intermediates as shown in Figure $3$. Figure $3$: Possible intermediates in the conversion of dinitrogen to ammonia by nitrogenase Nitrogenase can also other small molecules with triple bonds, including C=O: and H-C=C-H. The Structure of Nitrogenase Nitrogenase is a multiprotein complex in which the functional biological unit is built from two sets of the following dimeric structures: • a homodimer of subunits, E and F, which have binding sites for the mobile carrier of electrons (the protein ferredoxin or flavodoxin), ATP, and an FeS cofactor (4Fe-4S, called the F cluster) which accept electrons. These subunits are hence called the nitrogenase reductase subunits • a heterodimer of alpha and beta subunits. The a (alpha chain) binds the 8Fe-7S F cluster and the Fe-S-Mo M cluster. These subunits comprise the (di)nitrogenase catalytic subunits. The iron-molybdenum M cluster is in the α subunit and is where N2 reduction occurs. The P-cluster is between the α and β subunits and facilitates electron flow between the Fe-protein (F cluster) and FeMo-cofactor (M cluster) For clarity, one-half of the overall structure of the protein complex with bound ATP and metal centers is shown in Figure $4$. Figure $4$: Nitrogenase structure (4wzb) This half-structure consists of a homodimer of the reductase monomers and a heterodimer of nitrogenase subunits. Figure $5$ shows an interactive iCn3D model of the half structure of nitrogenase complex from Azotobacter vinelandii (4WZB) (long load). Figure $5$: Nitrogenase complex from Azotobacter vinelandii (4WZB). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...zPNjfPRwq8MSeA (long load)) The structure is color-coded in a fashion similar to Figure 4. The F cluster is labeled as SF4, the P cluster as CLF (FE(8)-S(7) cluster), and the M cluster as ICS (iron-sulfur-molybdenum cluster with interstitial carbon). The reductase subunits (called Av2 in Azotobacter vinelandii), accept electrons from ferredoxin and is where the ATP analog, ACP (phosphomethylphosphonic acid adenylate ester), and the 4Fe-4S F cluster is bound. The nitrogenase subunits (called Av1 in Azotobacter vinelandii) convert N2 to NH3 and is where the 8Fe-7S P cluster, FeMo (C Fe7 Mo S9) M Cluster is bound. An enhanced view of the bound cofactors and ATP is shown in the same spatial orientation in Figure $6$below. Figure $6$: Bound cofactor metal clusters and ATP in nitrogenase You can easily image the direction of the flow of electrons from the F cluster to the P cluster to the M cluster. The metal centers are shown in Figure $7$ in more detail in both line and space fill views. Figure $7$: Detailed structures of the metal cofactors in nitrogenase Mo is bound to 3 sulfur ions and oxygen from 3-hydroxy-3-carboxy-adipic acid as shown in Figure 7 above. The M cluster has an interstitial carbide ion that derives from -CH3 attached to the sulfur of S-adenosyl-methionine (SAM) allowing the carbide to be labeled with either 13C or 14C for mechanistic studies. These labeled carbides are not exchanged or used as a substrate when the enzyme undergoes catalytic turnover. Hence it seems that the carbide probably just stabilizes the M cluster. it won't be shown in the figures below showing more detailed mechanisms. We'll focus on two features of the reaction mechanism, ATP hydrolysis and the flow of electrons and protons to N2 as it is reduced to NH3. ATP hydrolysis ATP binds in the reductase subunit (AV2) where ferredoxin brings in electrons, and where the F metal cluster is bound. Much as GTP hydrolysis controls conformational change and subunit dissociation in the heterotrimeric Gαβγ in signal transductions, ATP hydrolysis in reductase (AV2) subunits, drives not only electron transfer but dissociation/reassociation of the reductase and nitrogenase catalytic subunits. It appears that 2 ATPs are hydrolyzed per electron transferred from the F cluster to the Mo M cluster. Since the oxidation numbers of N are 0 and -3 in N2 and NH3, respectively, sequential rounds of ATP hydrolysis and dissociation/reassociation occur.  Note that the Fe-protein hydrolyzes ATP only when bound to the MoFe-protein. The overall reaction involves the reduction of N2 to two molecules of NH3 at the FeMo cofactor.  It involves a reductive elimination of hydrides that are bridged by Fe ions (Fe-H-Fe) in a reaction that also makes H2 as a by-product.  We'll describe this complex reaction next. Nitrogenase Reaction: Part 1 - Addition of Electrons and Protons The sequential path of electrons from the reductase subunit containing the F cluster to the P and M clusters in the nitrogenase subunit should be apparent from the figures above. We will concentrate on the binding of N2 and how it receives electrons from the M cluster. Figure $8$ shows the FeMo-cofactor and some adjacent amino acid residues. Mo is labeled but not shown in spacefill. HCA is a bound molecule of 3-hydroxy-3-carboxy-adipic acid, which interacts with Mo. The carbide is shown in the middle as the green sphere and labeled CX. Figure $8$: Side chains near the M cluster If Val 70 is mutated to Ile, a substrate appears not to access the cluster suggesting that N2 may interact with the top part of the structure with the residues shown acting as gatekeepers. His side chains are often found at enzyme active sites so you might expect His 195 to be a general acid/base. Mutations lead to drastic losses in the reduction of N2. His 195 is involved in hydrogen bonds to sulfur S2B and bridges Fe2 and Fe3 in Figure 8, where reduction of N2 likely occurs. If His 195 moves, it can form short H bonds between the imidazole N and an H bond to HFe2. If the ring is rotated 1800, no proton transfer occurs from the surface. It appears that His195 might be involved in the first N2 protonation event. The Lowe and Thorneley (LT) model has been proposed as a mechanism for dinitrogen reduction. In this model, an electron and proton are added to the oxidized form of the enzyme (Eo) to produce E1. This is repeated 3 more times to form sequentially, E2, E3, and E4. Only then does N2 bind and the reduction of N2 occurs. Two of the added electrons are accepted by H+ ions which form H2, which is liberated on N2 binding. Hence only 6 electrons are added to the actual N2 molecule, in agreement with the change in oxidation numbers discussed above. The Lowe and Throneley model is shown in Figure $9$. Figure $9$: Lowe and Throneley model for electron and proton additions in nitrogenase The crystal structure shows 2 ATP analogs bound to the reductase subunit. The stoichiometry of the reaction shows 16 ATP used. Simple math suggests that 2 ATP are cleaved to support the entry of one electron into the complex, assuming 8 transferred electrons (6 to N2 and 2 to 2 protons to form H2). Part 1 - E1-E4: A potential structure for the E4 intermediate is shown in Figure $10$. Figure $10$: The E4 Janus intermediate in the reduction of N2. Note the central carbide is not shown. This is often called the Janus intermediate as it is halfway through the catalytic cycle. It is named for Janus, the Roman god of beginnings and transitions, and has been ascribed to gates, doors, doorways, and passages. Janus is typically shown with two faces, one looking to the future and one to the past (image below: DOI:10.1590/2177-6709.21.1.018-023.oin.  License CC BY 4.0 Creative Commons Attribution 4.0 International) The hydrides bridge 2 Fe ions so these are examples of three-center, two-electron bonds. The H+ ions in Figure 10 balance the charge from the hydrides. How does this reaction occur? We must look to organometallic chemistry to help us understand the mechanism of this and subsequent steps. Hydride equivalents have been added to the metals, associated with the oxidation of metal ions in the center. This particular reaction is called an oxidative addition. Presumably, the sulfur ions act as Lewis bases as they gain protons from a Lewis acid, probably His 195. Oxidative addition reactions Figure $11$ shows oxidative additions to metal centers for three different types of reactants. Figure $11$: Three different types of oxidative addition reactions (after Schaller, http://employees.csbsju.edu/cschaller/ROBI1.htm) In oxidative insertion, the oxidation state of the metal ion increases, hence the name oxidative. The hydrogens are now hydrides. Note the example for insertion of H2, an example similar to the proposed hydride additions to the M cluster. Oxidative reduction occurs most readily when the two oxidation states of the metal ion are stable. It is likewise favored for metal centers that are not sterically hindered (makes sense if A-B is to be added) and if A-B has a low bond dissociation energy. One way to study reaction intermediates is to trap them. If N2 can't access the binding site and the temperature is reduced, the accumulated hydrides (and for charge balance the H+s) in E4 might leave in the opposite reaction, reductive elimination, which we will discuss below, as the reaction goes back to E1. In the elimination, they could form H2 as metal gains back electrons in a reduction. The Val70Ile discussed above would allow an intermediate to be trapped. Nitrogenase Reaction: Part 2 - Reduction of N2 Step E4 to E5 seems a bit bizarre as H2 gas is released. This would seem to waste ATP but we should trust evolution has led to this mechanism for a reason. This mechanism, the reverse of oxidative addition, is another classic organometallic reaction, reductive elimination. Reductive elimination reactions In this reaction, a molecule is eliminated or expelled from the complex as the metal ion is reduced and adds two electrons. Figure $12$ shows reductive elimination. Reductive elimination occurs most readily in higher oxidation state metal centers which can be stabilized on reduction. It occurs most readily from electron-rich ligands and if the other surrounding ligands are bulky. The dissociating species must also be cis to each other in the transition metal complex so they can form a bond with each other when they leave. Figure $12$: Reductive Elimination reaction (after Schaller, ibid) Oxidative addition and reductive elimination (OA/RE) reactions at metal centers are often coupled together in organometallic catalytic cycles, in the same way as a histidine can act as a general acid and then accept a proton back as a general acid to complete the catalytic cycle. In the OA/RE reactions at metal centers, some rearrangements or other modifications can also occur. Think about it. The FeS clusters must return to their original oxidation state after the complete LT cycle. We will encounter another organometallic reaction after the addition of N2, migratory insertion, in the second half of the reaction. Another advantage of coupling OA/RE is that the positive charge or oxidation state on the transition metal complex does not get too high, which is unstable. Making a cation with a positive charge more positive becomes more difficult, much as removing a second proton from a polyprotic acid is more difficult than removing the first (as reflected in the higher pKa for removal of the second proton). Is H2(g) really released? To study this, investigators have used alternative substrates like acetylene, HC=CH (similar to N=N), in the presence of D2 and N2 in an aqueous system. It helps to see Figure $9$ again. Figure $9$: Lowe and Throneley model for electron and proton additions in nitrogenase The acetylene was reduced and formed C2H2D2 and C2H3D. Hence E4 must have had 2 Ds in it, and E2 probably 1. These results support the reversible reductive elimination mechanism for the E4 to E4:N2 reaction above. Previously it had been shown that H+ are reduced by D2 in the presence of D2 and N2 in an aqueous system, so these results are consistent. In additional support, deuterium from D2 is not incorporated into products (C2H2D2, C2H3D, or HD) in the absence of N2. Let's return for a moment to the bridging hydrides as shown again Figure $10$. Figure $10$: The E4 Janus intermediate in the reduction of N2 To summarize, it appears likely that the reductive elimination of the two proximal bridging hydrides is the mechanism for the formation of H2. The bridging hydrides, which are strong bases, are much less likely to be protonated than if the hydrides were terminal. A simple and competing protonation reaction could form H2 as well, and if that occurred the reducing equivalents of the bridging hydrides would be lost. Hence the bridging hydrides (share by two metal centers) are more stable and hence can "wait" for the incoming N2 reactants before their reducing equivalents are lost. They may convert to terminal hydrides eventually to facilitate substrate reduction (hydrogenation). The Janus intermediate E4 is now in a position to bind N2 and release H2. For each H- that binds to the M cluster, two H+s bind to the M cluster sulfides for electrostatic stabilization. Migratory Insertions We need to consider one last common type of reaction at a metal center, migratory insertions (MI). In a MI reaction, a group attached to a metal ion center is transferred to another group attached to the same metal. Figure $13$ shows four examples of MI reactions. Figure $13$: Migratory insertion reactions at metal centers Panel (a) shows a generic MI reaction. Panel (b) shows the interaction of carbon monoxide, :C=O: (isoelectronic and analogous to :N=N:), with a metal center. Panel (c) shows how a hydride could engage in 1:1 insertion as it shifts and covalently bonds to the first atom of another ligand bound to the metal center. Note that metal does not have to have a negative charge. This could theoretically be important for the reduction of N2 bound as a ligand through a coordinate covalent (dative) bond to the metal. Panel (d) shows the migration of an alkyl group. We will see that the MI reaction is involved in adding Hs to N2 starting not with N2 but with the N2H2 stage as the intermediates insert into an Fe-H bond. N2 is not reactive to the insertion of a hydride as is carbon monoxide, CO, which provides a positive oxygen to facilitate electron flow during the insertion. In addition, the oxygen become neutral after the reaction. This offers a great explanation for how Nature chose the FeMo cluster for nitrogen fixation. The interaction of two hydrides requires a 4 Fe face (coordinated with the carbon) that allows for the storage of reducing equivalents for the initial reduction of N2. The large M cluster is more stable and effectively held together by a central carbide anion. This also allows the metal centers to never change their oxidation state by more than 1 charge unit. The source of the protons to form N2H2 comes directly from the two H2 attached to the two sulfurs as they can't come from the hydrides which are released as H2. The seeming wasteful reductive elimination of H2 and energy is required to allow the kinetically unreactive N2 molecule to bind to the reduced and activate 4Fe face which is also electrostatically facilitated by the 2 bound protons in what has been called a push (reductive)-pull (protonation) reaction. This first step in N2 reduction to N2H2 is the hardest. Oxidation States of Nitrogenase Fe centers First Half: It would be difficult to assign specific oxidation states to each Fe ion in the M complex. Instead, we can assign relative changes in the oxidation states as the reaction proceeds from E0 to E4. In each step of the LT model, 1 electron is added. We will first assign this to an average Fe ion, M0, with an arbitrarily assigned oxidation state of 0. On the addition of 1 electron, the oxidation state would go from M0 to M-1 as the metal is reduced. The M-1 state is then oxidized as an electron is transferred to H+, and when 2 electrons are transferred, a single H- is made. The diagram in Figure $14$ shows the change in oxidation state in going from E0 to E4. Figure $14$: Change in oxidation state in going from E0 to E4 The red boxes highlight thermodynamic cycle-like steps which show how changes in the redox state of the Fe ions (M) could be visualized. Note that in going from E0 to E4, the actual oxidation state of M changes from 0 to +1 to 0 to +1 and back to 0. That is quite amazing given that 4 electrons have been added. Note also that in the red box going from step E0 to E1, M goes from -1 to +1 which corresponds to our description of an oxidative addition when the metal center loses two electrons. This mechanism shows that nitrogenase could be considered a "hydride storage device". Second Half (facing forward to production NH3): How does N2 initially interact with E4? It must depend on how the hydrides are released as H2, which evidence shows occurs by reductive elimination (re) and not hydride protonation (hp). On addition, the N2 very quickly is converted to diazene, HN=NH, with the departing H2 taking with it 2 H+s and 2 electrons (or reducing equivalents). These events could occur as shown in Figure $15$. Figure $15$: Reaction mechanism for the formation of N2H2 Now, with N2 bound as diazene (N2H2) and H2 released, the rest of the reaction could occur as shown below. One new step, a migratory insertion, is shown in Figure $16$. Figure $16$: Reaction mechanism for the conversion N2H2 to NH3. The two halves of the reaction are similar with bridging hydrides utilized - the E4 Janus intermediate links the two halves together.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/22%3A_Biosynthesis_of_Amino_Acids_Nucleotides_and_Related_Molecules/22.01%3A_Overview_of_Nitrogen_Metabolism.txt

Search Fundamentals of Biochemistry Introduction By the time many students get to the study of amino acid biosynthesis, they have seen so many pathways that learning new pathways for the amino acids seems daunting, even though they can be clustered into subpathways. Most know that from a nutrition perspective, amino acids can be divided into nonessential and essential (need external dietary supplementation) amino acids. These are shown for humans below. • Nonessential amino acids: Alanine, Asparagine, Aspartate, Cysteine, Glutamate, Glutamine, Glycine, Proline, Serine, Tyrosine • Essential amino acids: Arginine*, Histidine, Isoleucine, Leucine, Lysine, Methionine*, Phenylalanine*, Threonine, Tryptophan, Valine Three of the essential amino acids can be made in humans but need significant supplementation. Arginine is depleted in processing through the urea cycle. When cysteine is low, methionine is used to replace it so its levels fall. If tyrosine is low, phenylalanine is used to replace it. The amino acids can be synthesized from glycolytic and citric acid cycle intermediates as shown in Figure \(1\) Figure \(1\): Summary amino acid synthesis from glycolytic and TCA intermediates For this chapter subsection, we will provide only the basic synthetic pathways in abbreviated form without going into mechanistic or structural details Amino acid synthesis from glycolytic intermediates From Glucose-6-Phosphate: Histidine The synthesis of histidine from a phosphorylated form of ribose (derived from glucose-6-phosphate) is shown in Figure \(2\). Figure \(2\): Synthesis of histidine from a phosphorylated form of ribose From 3-phosphoglycerate: Serine, Glycine, and Cysteine The synthesis of serine, glycine, and cysteine from 3-phosphoglycerate is shown in Figure \(3\). Figure \(3\): The synthesis of serine, glycine, and cysteine from 3-phosphoglycerate From Phosphenol Pyruvate: The Aromatics - Trp, Phe, and Tyr The synthesis of the first of the biosynthetic pathways for the aromatic amino acids phenylalanine, tryptophan, and tyrosine from phosphoenolpyruvate up to chorismate is shown in Figure \(4\). Figure \(4\): Synthesis of the first of the biosynthetic pathways for the aromatic amino acids phenylalanine, tryptophan, and tyrosine from phosphoenolpyruvate up to chorismate Chorismate to tryptophan The synthesis of the second half of the biosynthetic pathway for tryptophan from chorismate is shown in Figure \(5\) Figure (5\): Synthesis of the second half of the biosynthetic pathways for the aromatic amino acid tryptophan from chorismate Chorismate to Phe and Tyr The synthesis of the second half of the biosynthetic pathway for phenylalanine and tyrosine from chorismate is shown in Figure \(6\) Figure \(6\): Synthesis of the second half of the biosynthetic pathway for phenylalanine and tyrosine from chorismate From Pyruvate: Ala, Val, Leu, Ile Ala can easily be synthesized from the alpha-keto acid pyruvate by a transamination reaction, so we will focus our attention on the others, the branched-chain nonpolar amino acids Val, Leu, and Ile. The synthesis of valine, leucine, and isoleucine from pyruvate is shown in Figure \(7\). Figure \(7\): The synthesis of valine, leucine, and isoleucine from pyruvate TCA Intermediates From α-ketogluatarate: Glu, Gln, Pro, Arg Since amino acid metabolism is so complex, it's important to constantly review past learning. Figure \(8\) from section 18.2 shows the relationship among Glu, Gln, and keto acids. Figure \(8\): Glutamate and glutamine synthesis from α-ketoglutarate As is evident from the figure, glutamic acid can be made directly through the transamination of α-ketoglutarate by an ammonia donor, while glutamine can be made by the action of glutamine synthase on glutamic acid. Arginine is synthesized in the urea cycle as we have seen before. It can be made from α-ketoglutarate through the following sequential intermediates: N-acetylglutamate, N-acetylglutamate-phosphate, N-acetylglutamate-semialdehyde, N-acetylornithine to N-acetylcitruline. The is deacetylated and enters the urea cycle. The pathway for conversion of α-ketoglutarate to proline is shown in Figure \(9\). Figure \(9\): Conversion of α-ketoglutarate to proline From oxalacetate: Asp, Asn, Met, Thr, Lys OAA to Aspartatic Acid This is a simple transamination Aspartic Acid to Asparagine This is catalyzed by the enzyme Asparagine Synthase as shown in the reaction equation below: Aspartate + Glutamine + ATP + H2O → Asparagine + Glutamic Acids + AMP + PPi Aspartic Acid to Lysine There are two pathways. • The diaminopimelic acid (DAP) pathway uses aspartate and pyruvate and forms diaminopimelic acid as an intermediate. It's found in bacteria, some fungi, and archaea and in plants. • The aminoadipic acid (AAA) pathway uses α-ketoglutarate and acetyl-CoA and forms aminoadipic acid as an intermediate. It is used by fungi., Here we present just the synthesis of lysine from aspartate and pyruvate using the diaminopimelic acid DAP pathway. The pathway is shown in Figure \(10\). Figure \(10\): The synthesis of lysine from aspartic acid in the diaminopimelic acid DAP pathway . Aspartic acid to Threonine The conversion of aspartic acid to threonine is shown in Figure \(11\). Figure \(11\): The conversion of aspartic acid to threonine Aspartic acid to Methionine The conversion of aspartic acid to methionine is shown in Figure \(12\). Figure \(12\): The conversion of aspartic acid to methionine

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/22%3A_Biosynthesis_of_Amino_Acids_Nucleotides_and_Related_Molecules/22.02%3A_Biosynthesis_of_Amino_Acids.txt

Search Fundamentals of Biochemistry Introduction Once made or ingested, amino acids have many metabolic fates. Of course, they are used for the synthesis of proteins. Aspartate and glutamate (and indirectly glutamine) can be converted to oxaloacetate and α-ketoglutarate, respectively, and used in the citric acid cycle for energy production. They can also be used for gluconeogenesis using mitochondrial and cytoplasmic enzymes. Branched-chain amino acids can be converted to acetyl-CoA and used in energy production or fat synthesis. A review summary of the use of amino acids in energy and biosynthetic metabolic pathways is shown in Figure \(1\). Figure \(1\): Review summary of the use of amino acid in energy and biosynthetic metabolic pathways. Lieu, E.L., Nguyen, T., Rhyne, S., et al. Amino acids in cancer. Exp Mol Med 52, 15–30 (2020). https://doi.org/10.1038/s12276-020-0375-3. Creative Commons Attribution 4.0 International License, http://creativecommons.org/licenses/by/4.0/. Amino acids are shown in green and other metabolites are in red. Orange represents transporters. Yellow boxes signify enzymes. Lesser known abbreviations for species include SHMT1 serine hydroxymethyltransferase, cytosolic, BCAT branched-chain amino acid transaminase, mitochondrial, BCAA branched-chain amino acid (valine, leucine, isoleucine), BCKA branched-chain ketoacid, GOT1 aspartate transaminase, cytosolic (AST), GLS glutaminase, GS glutamine synthetase (cytosolic and mitochondrial), ASNS asparagine synthetase, PRODH pyrroline-5-carboxylate dehydrogenase, PYCR pyrroline-5-carboxylate reductase, P5C pyrroline-5-carboxylate, GSH glutathione, PRPP phosphoribosyl pyrophosphate, LAT1 large-neutral amino acid transporter 1, SLC25A44 solute carrier family 25 member 44, GLUT glucose transporter, Cancer cells from an increased need for fuels and biosynthetic intermediates. Both can come from amino acids as described previously. Glutamine is a key amino acid, especially if glucose is depleted as α-ketoglutarate (α-KG) and subsequently oxaloacetate (OAA generated from it powers the TCA cycle as fumarate, malate, and citrate are significantly increased. Hence it is both anaplerotic and a source of fuel. Similar increases in citrate occur in hypoxia. Aerobic glycolysis (Warburg effect) occurs in cancer cells, which show enhanced glucose uptake and conversion to lactate even in the presence of oxygen. This process can go so quickly that the amount of ATP produced in cancer cells from aerobic glycolysis can be similar to the from oxidative metabolism in the mitochondria, even though it is far less efficient. More information on cancer cell metabolism is found in Chapter 23. In this chapter, we will discuss the conversion of amino acids to other molecules not directly involved in those metabolic pathways. We will focus on their use for the synthesis of polyamines, heme, and neurotransmitters in this chapter section. We won't discuss detailed mechanisms or structures for the proteins and enzymes involved in these pathways. In the next chapter section (22.4), we will present amino acids as substrates in the synthesis of pyrimidine and purine bases for nucleotides and nucleic acids. Polyamine synthesis If a non-quaternary amine has a single positive charge when protonated, a polyamine can have multiple positive charges. Hence they would be expected to bind to almost any negatively charged biomolecule, but especially those with multiple negative charges. These would include the polyanions RNA and DNA as well as proteins and lipid bilayers. They would then have the potential ability to regulate many features of cell life, including DNA replication and transcription, RNA translation, and a multitude of binding interactions. The question arises if these interactions are nonspecific, or specific, in which case they can be considered key regulators of cellular activity. Polyamine response elements have been found that regulate the transcription of genes including c-Myc and c-Jun. Polyamines have been shown tumor growth and aggressiveness. The main biological polyamines include putrescine, spermine, and spermidine. Another is cadaverine. Given their names, you can surmise that they smell horrible. The synthesis of three polyamines from arginine and SAM is shown in Figure \(2\). Figure \(2\): Polyamine synthesis form arginine and SAM Glutathione synthesis and redox balance Glutathione, γ-glutamylcysteinylglycine (GSH), is a chief regulator of the oxidation state of a cell. As a disulfide bond can be cleaved and hence reduced by the excess concentration of a thiol (sulfhydryl) like b-mercaptoethanol (which gets oxidized in the process), the free thiol on glutathione can act as a reducing agent in the cell. The production of reactive oxygen species (ROS) in normal but especially tumor cells, which have increased O2 demand and use, is countered by the generation of an antioxidant defense state. This is characterized in part by increased levels of reductants such as NADPH but especially glutathione. It can react with H2O2 through the enzyme GSH peroxidase to form water and the oxidized disulfide form of GSH, GSSG. The GSSG is oxide back to GSH by glutathione reductase (GR) and NADPH. Figure \(3\) shows the synthesis of glutathione from glutamate, cysteine, and glycine. Figure \(3\): Synthesis of glutathione NADPH is generated in the cell by the phosphopentose pathway metabolism of glucose and by malic enzyme. It can also be generated from the Ser-Glycine One Carbon Cycle (SGOT) that we saw in Chapter 18.4, which is shown again in Figure \(4\). Under appropriate conditions, this cycle can produce NADPH. Figure \(4\): The Ser-Gly One Carbon (SGOC) Cycle Serine Hydroxymethyltransferases (SHMTs) 2 is upregulated by HIF1α and helps maintain the NADPH/NADP+ ratio. Given the connection between the SGOC and the methionine cycle through folate, a decrease in serine concentration leads to a decrease in GSH. Heme Biosynthesis This section is derived from Aminat S. Ogun; Neena V. Joy; Menogh Valentine. https://www.ncbi.nlm.nih.gov/books/NBK537329/. Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/) Heme is a macrocyclic tetrapyrrole ring structure containing two nonpolar vinyl groups on one edge and two charge propionates on the other. It is extensively conjugated with 26 π electrons (4n+2 = 4(6)+2) so it is aromatic. The molecule without Fe2+ is called protoporphyrin IX and with a centrally-coordinate Fe2+, it is called heme. The structures of both are shown in Figure \(5\). Figure \(5\): Structure of protoporphyrin IX and heme It is found in oxygen-binding proteins and as substrates and cofactors for enzymes involved in electron transport. It is synthesized in the bone marrow and liver. Alternative forms of heme include heme b (in hemoglobin), heme a (cytochrome a), and heme c (cytochrome c). Its synthesis, as expected given its macrocyclic structure, is complicated. The key enzyme in the pathway for regulation is 5'-Aminolevulinic acid synthase (ALA-S). Liver and bone express ALAS2 while ALAS1 is expressed in all tissues. The synthesis starts in the mitochondria and ends in the cytosol. The overall pathway for heme synthesis is shown in Figure \(6\). Figure \(6\): Heme biosynthetic pathway. Wikimedia Commonsile: Heme-Synthesis-Chemical-Details-Mirror.svg 5'-Aminolevulinic acid synthase (ALA-S), a pyridoxal phosphate-dependent enzyme, catalyzes the rate-limiting step in heme synthesis in the liver and erythroid cells. It is highly regulated There are two forms of ALA Synthase, ALAS1, and ALAS2. All cells express ALAS1 while only the liver and bone marrow expresses ALAS2. The gene for ALAS2 is on the X-chromosome. After the synthesis of ALA in the mitochondria, it moves into the cytoplasm for the remaining steps. Figure \(7\) shows a likely mechanism for the first committed step, the production of ALA. This enzyme is used in the synthesis of all tetrapyrroles, including heme, chlorophyll, and cobalamin. Figure \(7\): Mechanism for 5'-Aminolevulinic acid synthesis by ALAS (Wikipedia. https://en.Wikipedia.org/wiki/Aminol..._acid_synthase) The pathway shown in Figure 6 above is called the C4 pathway and is found in mammals, fungi, and purple nonsulfur bacteria. A C5 pathway is found in most bacteria, all archaea, and plants. The biosynthetic pathway for heme synthesis in E. Coli is shown in Figure \(8\). Figure \(8\): Heme pathway in E. coli. Zhang, J., Kang, Z., Chen, J. et al. Optimization of the heme biosynthesis pathway for the production of 5-aminolevulinic acid in Escherichia coli. Sci Rep 5, 8584 (2015). https://doi.org/10.1038/srep08584. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ The pathway is divided into three modules (module I, module II, and module III in the dotted box). The arrows in green and red represent the enzymes that are positive and negative to ALA accumulation, respectively. Dotted red arrows represent the feedback inhibition. α-KG: α-ketoglutarate, GSA: glutamate-1-semialdehyde, ALA: 5-aminolevulinic acid, PBG: porphobilinogen, HMB: hydroxymethylbilane, GltX: glutamyl-tRNA synthetase, HemA: glutamyl-tRNA reductase, HemL: glutamate-1-semialdehyde aminotransferase, HemB: 5-aminolevulinic acid dehydratase, HemC: porphobilinogen deaminase, HemD: uroporphyrinogen III synthase, HemE: uroporphyrinogen decarboxylase, HemF: coproporphyrinogen III oxidase, HemG: protoporphyrin oxidase, HemH: ferrochelatase. In immature red blood cells (reticulocytes), heme increase globin protein synthesis. The hormone erythropoietin increases heme synthesis. In the liver, heme is part of cytochrome P450s. Increased concentration of drugs causes increases in ALAS1 to produce the cytochrome P450s to metabolize them. Also, low heme concentration increases ALAS1 transcription. Mutations in ALAS2 can lead to X-linked sideroblastic anemia from decreased heme production even as Fe2+ continues to enter the cell. Yeast ALAS is a homodimer with PLP covalently attached through a Schiff base link to lysine 337 of just one of the monomers. The structures of a noncovalent complex of PLP with ALAS (pdb 5TXR) and the covalently bound one (5TXT) show large changes in the protein conformation. PLP when covalently attached reorders the active. A C-terminal extension not found in bacteria wraps around the dimer and binds near the active site and is important for activity. Mutations in the tail can result in human diseases. Figure \(9\) shows an interactive iCn3D model of the 5-aminolevulinic acid synthase with covalently attached PLP (5TXT). Figure \(9\): 5-aminolevulinic acid synthase with covalently attached PLP (5TXT). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?fTBbWuS3HPTP8uRm9 Lysine 337 (spacefill, CPK colors, labeled) in the A chain (magenta, no bound PLP) is shown. Lys 337 in the B chain (cyan) is covalently linked to PLP. The side chain of lysine 337 covalently attached to PLP is shown in spacefill, CPK colors, and labeled. The C-terminal extension (493–548) is shown as a red backbone chain. The very distal end of the extension is disordered and missing in the B chain (cyan). Figure \(10\) shows an interactive iCn3D model of aligned 5-aminolevulinic acid synthase with free PLP (not covalently attached, 5TXR) and with covalently attached PLP (5TXT ). Figure \(11\): Alignment of 5-aminolevulinic acid synthase with free PLP (5TXR) and with and with covalently attached PLP (5TXT ). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...aqN4Qpi2pSLkD7 The 5TXT structure contains two molecules of a stabilizing molecule shown in stick form, which you can ignore. The A chain is shown in magenta and the B chain is in cyan. Press "a" to toggle back and forth between the structures. The C-terminal extension is missing from the figure. Figure \(11\) shows another view of heme synthesis which emphasizes the role of mitochondrial and cytoplasmic enzymes. Neurotransmitters The section below is modified from Manorama Patri. Synaptic Transmission and Amino Acid Neurotransmitters. DOI: 10.5772/intechopen.82121. https://www.intechopen.com/books/neu...rotransmitters. Creative Commons Attribution 3.0 License, There are three major categories of amino acids and their derivatives act as neurotransmitters are: 1. Amino acids: The neurotransmitters of this group are involved in fast synaptic transmission and are inhibitory and excitatory in action (primarily glutamic acid, GABA, aspartic acid, and glycine). 2. Amines: Amines are modified amino acids such as biogenic amines, e.g., catecholamines. The neurotransmitters of this group involve in slow synaptic transmission and are inhibitory and excitatory in action (noradrenaline, adrenaline, dopamine, serotonin, and histamine). 3. Others: The ones which do not fit in any of these categories (acetylcholine and nitric oxide). Amino acids are among the most abundant of all neurotransmitters present within the central nervous system (CNS). Amino acid transmitters provide the majority of excitatory and inhibitory neurotransmission in the nervous system. Amino acids used for synaptic transmission are compartmentalized (e.g., glutamate, compartmentalized from metabolic glutamate used for protein synthesis by packaging the transmitter into synaptic vesicles for subsequent Ca2+-dependent release). Amino acid neurotransmitters are all products of intermediary metabolism except GABA. Unlike all the other amino acid neurotransmitters, GABA is not used in protein synthesis and is produced by an enzyme (glutamic acid decarboxylase; GAD) uniquely located in neurons. Here is some more specific information: • Glutamate: Glutamate is used at the great majority of fast excitatory synapses in the brain and spinal cord. Glutamate binds to glutamate receptors of which there are many subtypes based on other molecules (some amino acid derivatives) that can bind to them. These other molecules include N-methyl-D-aspartate (NMDA), α-amino-3-hydroxy-5-methyl-4-isoxazolepropionate (AMPA) kainate, and quisqualate. • Aspartate: Aspartate is the most abundant excitatory neurotransmitter in the CNS. Like glycine, aspartate is primarily localized to the ventral spinal cord. Note that the two major excitatory neurotransmitters both have carboxylic acid side chains. • Gamma-aminobutyric acid (GABA): GABA, which is not one of the canonical amino acids used in protein biosynthesis, is the most ubiquitous inhibitory neurotransmitter in the brain. • Glycine: lycine receptors are ligand-gated ion channels that increase Cl influx and hence are generally inhibitory. Hydroxymethyl transferase converts the amino acid serine to glycine. Glycine has been found to play a role in the functional modulation of NMDA receptors The pathways for the synthesis of amino acid-derived bioactive amines and neurotransmitters are shown in Figure \(12\). Figure \(12\): Pathways for the synthesis of amino acid-derived bioactive amines and neurotransmitters Note the structural similarity of the psychotropic and hallucinogenic drug LSD to serotonin (5HT), amphetamines to norepinephrine and epinephrine, and melatonin (a substance some take as a sleeping aid and which forms in the dark at night in brains. The name catecholamines derive from the common name of the 1,2-dihydroxybenzene group (catechol). The first and rate-limiting step in catecholamine synthesis is catalyzed by tyrosine hydroxylase (TH). It has no heme but it has an Fe2+ and tetrahydrobiopterin as a cofactor used in the synthesis of dihydroxyphenylalanine (DOPA). Tyrosine hydroxylase is rate-limiting for the synthesis of all three transmitters. The enzyme is inhibited by catecholamines including dopamine, a downstream product, and is activated by phosphorylation on serine 40. The structures of TH in the absence of dopamine and the pSer40 state are known. The protein is a tetramer with a regulatory domain (dimer) and catalytic domain (also a dimer) separated by 15 Å. The mammalian TH is a member of the aromatic amino acid hydroxylases (AAAHs) which are mainly found as homotetramers. Each subunit has 3 domains: • The N-terminal regulatory domain (RD) that has an unstructured variable-length section followed by an ACT (aspartate kinase-chorismate mutase-TyrA) domain. The N-terminal tail contains serine 40 which on phosphorylation relieves the inhibition when dopamine is bound to the catalytic domain. • a central catalytic domain (CD) that binds Fe2+, aromatic amino acid substrates, and the tetrahydrobiopterin cofactor • C-terminal oligomerization domain (OD) which leads to dimer and tetramer formation. Figure \(13\) shows a potential model that illustrates dopamine (DA)-mediated feedback inhibition and its regulation by serine 40 phosphorylation through the interaction of the N-terminal tail of the regulatory domain (RD) with the catalytic domain. All forms containing bound dopamine (yellow star) are inactive. Figure \(13\): Cartoon model of DA-mediated feedback inhibition and its regulation by S40 phosphorylation. Bueno-Carrasco, M.T., Cuéllar, J., Flydal, M.I. et al. Structural mechanism for tyrosine hydroxylase inhibition by dopamine and reactivation by Ser40 phosphorylation. Nat Commun 13, 74 (2022). https://doi.org/10.1038/s41467-021-27657-y. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/. In the active, apo, and non-phospho states, the 39−58 α-helix of the N-terminal regulatory domain of TH is detached from the main structure (I, apo-TH). The feedback inhibitor DA binds to the TH active site, most likely in the open conformation (I′, TH(DA)). DA-binding favors the interaction of the N-terminal α-helix with the same binding site, which blocks DA exit and contributes to the high-affinity binding and strong inhibition of TH activity (II, TH(DA)). Protein Kinase (PK) phosphorylation of S40 in TH(DA), leads to state III (THS40p(DA)), prompting the detachment of the α-helix from the TH active site (IV′), which opens up for DA-dissociation and activation (IV, THS40p). PKs and protein phosphatase(s) (PP) control the transition between THS40p and unphosphorylated TH for both DA bound (I′ ↔ IV′ and II ↔ III) and apo-TH (I ↔ IV). States I′ and III are expected to be only transiently populated during DA binding as states II and IV' will be more stable. Hence states I′ and III states are faded. S40 is also expected to be less accessible in state II than in state I, which is indicated by stippled lines for phosphorylation of TH in state II. The case for dephosphorylation is not known, but it could be expected that state III is a poorer substrate for PP than the open states IV′ and IV. The dephosphorylation reaction III → II is therefore also stippled. The states where we provide structural details in this work (I, II, and IV) are marked with circles. Figure \(14\) shows a model of the TH active site changes on phosphorylation of serine 40 using structural and molecular dynamics approaches that led to the cartoon model above. Figure \(14\): Modeling of the TH active site. Panel (a) shows models demonstrating the effect of serine 40 phosphorylation on the interaction of the N-terminal α-helix with bound dopamine (DA). Representative conformations from the last 50 ns of a 500 ns MD simulations for TH(DA) (grey ribbon) and pS40-TH(DA) (light blue ribbon) are shown. The resulting structures show a slight shift of the N-terminal α-helix upon phosphorylation, most probably due to electrostatic repulsion between the phosphate and E325, E375, and D424. Panel (b) shows a detailed view of the atomic model of the TH(DA) active site. (left) The N-terminal α-helix (orange), establishes connections with the adjacent helix D360-E375 and with residues of the 290–297 and 420–429 loops (blue, right). Pane (c) shows a cartoon depicting the interactions established between residues of the N-terminal α-helix that enters the active site, and residues of adjacent regions. Figure \(\PageIndex{15\) below shows an interactive iCn3D model of the full-length tyrosine hydroxylase in complex with dopamine (residues 40-497) in which the regulatory domain (residues 40-165) has been included only with the backbone atoms (6zvp) Figure \(15\): Tyrosine hydroxylase dopamine complex (6zvp). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...hu6vJn22eDkWw7 Finally, Figure \(16\) shows an interactive iCn3D model of the active site of tyrosine hydroxylase in complex with dopamine (6zvp) Figure \(16\): Active site of tyrosine hydroxylase dopamine complex (6zvp). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?sx5obQnzojmKXuLS9 The side chains binding the active site Fe2+ and the interaction of Fe2+ with dopamine (LDP) are shown in sticks and labeled. The oxygen of serine 40 is shown as a red sphere. Dopamine is the molecule containing the 1,2-dihydroxybenzene. Figure \(17\) shows a final summary presentation of the conversion of phenylalanine, tyrosine, and tryptophan to neurotransmitters. Figure \(17\): Comparison of monoamine synthesis pathways. Adapted from Hochman, Shawn. (2015). Neural Regeneration Research. 10. 10.4103/1673-5374.169625. 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textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/22%3A_Biosynthesis_of_Amino_Acids_Nucleotides_and_Related_Molecules/22.03%3A_Molecules_Derived_from_Amino_Acids.txt

Search Fundamentals of Biochemistry Introduction We conclude our exploration of metabolic pathways with the biosynthesis and breakdown of nucleotides, the monomers that comprise nucleic acids. We can't also forget the important role of ATP as the universal carrier of biological free energy, as well as the nucleotides involved in signal transduction (GTP in heterotrimeric G proteins, small G proteins, and ATP as substrate in protein phosphorylations by kinases). As with the other later sections on metabolism, we won't focus much on detailed reaction mechanisms or enzyme structures, with one exception, the enzyme that converts nucleotides to deoxynucleotides. Nucleotide synthesis is often included in chapters on amino acid metabolism as almost every atom in the purine and pyrimidine ring derives from them as shown in Figure \(1\). Figure \(1\): Source of atoms in nucleotide bases. Lieu, E.L., Nguyen, T., Rhyne, S., et al. Amino acids in cancer. Exp Mol Med 52, 15–30 (2020). https://doi.org/10.1038/s12276-020-0375-3. Creative Commons Attribution 4.0 International License, http://creativecommons.org/licenses/by/4.0/. For purines, glutamine and aspartate provide the nitrogen for the nucleotide's rings. They also provide the NH3s for the ring substituents (glutamine for adenine and aspartate for guanine). Glycine and formate provide the carbon atoms for the rings. The one carbon molecule formate, which derives from glycine, is also added to purine rings. Glycine provides a one-carbon unit indirectly through the main carrier of activated one-carbon units, 5,10-meTHF, which is converted to formate through 10-formyl THF. Pyrimidines are much smaller and their synthetic pathway reflects that. Instead of being synthesized as nucleobases as in the case of purines, they are made as ribonucleotides as they are linked to phosphoribosyl pyrophosphate (PRPP). Glutamine and aspartate again provide the ring C and N atoms. The one-carbon unit derives from serine-to-glycine conversion. A methyl group from the activated 1C donor, 5,10-meTHF, is added to dUMP to make dTMP. Purine Synthesis The material below derives from De Vitto, H.; Arachchige, D.B.; Richardson, B.C.; French, J.B. The Intersection of Purine and Mitochondrial Metabolism in Cancer. Cells 2021, 10, 2603. https://doi.org/10.3390/cells10102603. Creative Commons Attribution License Mammals have two pathways for purine synthesis, a de novo pathway and a salvage pathway to recycle nucleotide bases. The salvage pathway is typically sufficient as purine bases come from nucleic acid breakdown. The resulting free bases (adenine, guanine, and hypoxanthine are connected to phosphoribosyl pyrophosphate (PRPP) to form nucleoside monophosphates (NMP) using either adenine phosphoribosyltransferase (APRT) to form AMP or hypoxanthine-guanine phosphoribosyltransferase (HGRT) to form IMP and GMP. PRPP is a substrate in both the salvage and de novo pathways. The overall de novo and salvage pathways for purine synthesis are described in detail in Figure \(2\). Figure \(2\): Purine metabolic pathways. De Vitto, H.; Arachchige, D.B.; Richardson, B.C.; French, J.B. The Intersection of Purine and Mitochondrial Metabolism in Cancer. Cells 2021, 10, 2603. https://doi.org/10.3390/cells10102603. Creative Commons Attribution License The conserved de novo biosynthesis pathway to generate IMP consists of 10 chemical steps catalyzed by 6 gene products in humans. These include the trifunctional enzyme TGART, composed of GAR synthetase (GARS), GAR transformylase (GARTfase), and AIR synthetase (AIRS) domains; the bifunctional enzymes PAICS, composed of CAIR synthetase/AIR carboxylase (CAIRS) and SAICAR synthetase (SAICARS), and ATIC, composed of AICAR transformylase (AICART) and IMP cyclohydrolase (IMPCH); and three monofunctional enzymes, phosphoribosyl amidotransferase (PPAT), formyl glycin amidine ribonucleotide synthetase (FGAMS), and adenylosuccinate lyase (ADSL). Downstream IMP is converted to (1) GMP through stepwise reactions of IMP dehydrogenase (IMPDH) followed by GMP synthetase (GMPS) and (2) AMP via adenylosuccinate synthetase (ADSS) followed by ADSL. The salvage pathway requires PRPP to generate IMP and GMP through one-step reactions mediated by hypoxanthine phosphoribosyltransferase (HPRT) utilizing hypoxanthine and guanine bases. AMP is generated by adenine phosphoribosyltransferase (APRT) utilizing adenine base and PRPP as substrates. Mitochondria supply precursors for purine de novo biosynthesis including glycine, N10-formyl THF, and aspartic acid through their one-carbon cycle (1C cycle) and tricarboxylic acid cycle (TCA). The de novo pathway kicks in when there is high demand for purines. Six enzymes are required for the 10-step pathway. Three of these are multifunctional enzymes catalyzing multiple steps in the pathway, comprising the two bifunctional enzymes phosphoribosylaminoimidazole carboxylase (PAICS) and AICAR transformylase/inosine monophosphate cyclohydrolase (ATIC) and the trifunctional enzyme glycinamide ribonucleotide transformylase (TGART). When active, the pathway is limited both by substrate availability and by the reaction rate of its initial step, the conversion of PRPP to phosphoribosylamine (PRA) by phosphoribosylpyrophosphate amidotransferase (PPAT). The final product of the de novo biosynthesis pathway, IMP is a substrate for the production of both AMP and GMP. 6 ATP are used to make 1 IMP from PRPP. None are required in the salvage pathway. PPAT is also called Glutamine phosphoribosylpyrophosphate amidotransferase or amidophosphoribosyltransferase.  It catalyzes the rate limiting step, is tightly regulated. PPAT possesses two nucleotide-binding sites near the active site, allowing for feedback control by downstream purine nucleotides via allosteric inhibition. Figure \(3\) shows an interactive iCn3D model of the Glutamine Phosphoribosylpyrophosphate Amidotransferase from Arabidopsis thaliana (6LBP) Figure \(3\): Glutamine Phosphoribosylpyrophosphate Amidotransferase from Arabidopsis thaliana (6LBP). (Copyright; author via source). Click the image for a popup or use this external link:  https://structure.ncbi.nlm.nih.gov/i...uWnf1bTq1odk88 The plant enzyme is a homotetramer, which each subunit having a Fe4S4 center (spacefill).  The active site residues in each subunit (DDS 432-444 and DS 369-370) are shown as colored sticks and labeled. Regulation of IMP production also occurs through enzyme phosphorylation. For example, Thr 397 on PPAT is phosphorylated by protein kinase B (PKB). PRPP concentrations also affects the rate. Another regulation of the flux through the de novo pathway is through the condensation of the enzymes into the purinosome, which contains PPAT, TGART, formylglycinamidine ribonucleotide synthetase (FGAMS), PAICS, adenylosuccinate lyase (ADSL), and ATIC. The purinosome also interacts with the mitochondria which would allow high local concentrations of ATP. A cartoon view of the purinosome is shown in Figure \(4\). Figure \(4\): Cartoon showing the purinosome. Baresova et al (2018) PLoS ONE 13(7): e0201432. https://doi.org/10.1371/journal.pone.0201432. Creative Commons Attribution License, Figure \(5\) shows another view of de novo IMP synthesis in which the origin of each atom in the purine ring is shown in color. Figure \(5\): De novo IMP synthesis showing the origin of atoms in IMP. Figure \(6\) shows an expanded view of the conversion of IMP to GTP and ATP. Figure \(6\): Conversion of IMP to GTP and ATP Pyrimidine Synthesis As mentioned in the introduction, pyrimidines have a much simpler biosynthetic pathway. Instead of being synthesized as nucleobases as in the case for purines, they are made as ribonucleotides as they are linked to phosphoribosyl pyrophosphate (PRPP). Glutamine and aspartate again provide the ring C and N atoms. The one-carbon unit derives from serine-to-glycine conversion. A methyl group from the activated 1C donor, 5,10-meTHF, is added to dUMP to make dTMP. The first step in the pathway for pyrimidine synthesis is the condensation of aspartate and carbamoyl phosphate. We have seen the synthesis of carbamoyl phosphate in the urea cycle by the enzyme carbamoylphosphate synthase I (CPSI) in Chapter 18.3 but present the reaction again in Figure \(7\). Figure \(7\): Synthesis of carbamoyl phosphate A different cytosolic version of the enzyme, CPS II, is used to synthesize both arginine and pyrimidine nucleotides. It uses glutamine as a donor of NH3. The pathway for the synthesis of UTP and CTP are shown in Figure \(7\). It does not explicitly show the synthesis of carbamoylphosphate, which is an intergyral part of the pathway and one of the rate-limiting steps in pyrimidine synthesis. Figure \(8\): De novo synthesis of UDP, UTP and CTP UDP and CDP can be converted to dCDP and dUDP, then on to dCPT and dUTP, and to dTMP as shown in Figure \(9\). Figure \(9\): Synthesis of dTMP Some of the material below derives from Li et al. Int. J. Mol. Sci. 2021, 22(19), 10253; https://doi.org/10.3390/ijms221910253. Creative Commons Attribution (CC BY) license (https://creativecommons.org/licenses/by/4.0/) In purine synthesis, we saw multifunctional enzymes that catalyze several steps, as well as the assembly of the enzymes in the de novo pathway into the purinosome. In an analogous fashion, three different enzyme activities that catalyze the first three combined rate limiting steps of pyrimidine synthesis, Carbamoyl-phosphate synthetase, Aspartate transcarbamoylase, and Dihydroorotase are found in a single, multifunction protein referred to as CAD. Its structure is a hexamer of a 243K monomer. It has 4 domains that include • glutamine amidotransferase (GATase) which "moves" HCO3, glutamine, and ATP to the CPSIIase domain • carbamoylphosphate synthetase II (CPSIIase): This has two parts, CPSaseA, and CPSase B. They combine functionally with GATase to form a glutamine-dependent carbamoylphosphate synthase (CPSase) • aspartate transcarbamylase (ATCase) acts as a homotrimer • dihydroorotase (DHOase) catalyzes the reversible cyclization reaction. The CAD protein is a 'fusion' protein encoding these four enzymatic activities of the pyrimidine pathway. Figure \(10\) shows the domain structure of the CAD protein. Figure \(10\): Domain structure of CAD The red represents glutamine amidotransferase and the blue the carbamoyl phosphate synthase ATP binding domain. More specifically, the following amino acid stretches comprise the different domains: GATase (2-365), CPSase A (395-933), CPSlase B (934-1455), DHOase (1456-1788), and ATCase (1918-2225) Figure \(11\) shows an interactive iCn3D model of the AlphaFold predicted model of the CAD protein (P27708) Figure \(11\): AlphaFold predicted model of the CAD protein (P27708). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...5V2XUmfsz1qjZ6 The GATase domain is magenta, the CPSase domains orange, the DHOase domain yellow, and the ATCase domain cyan. The structure of the disordered loops could not be modeled. Ribonucleotide reductases (RNRs) Ruskoski and. Boal. J. Biol. Chem. (2021) 297(4) 101137. DOI:https://doi.org/10.1016/j.jbc.2021.101137. CC-BY license (http://creativecommons.org/licenses/by/4.0/). Ribonucleotide reductases (RNRs), also called ribonucleoside-diphosphate reductase, catalyze the oxidation of the C2'-OH on the ribose ring to C2'-H through a free radical mechanism for the oxidation of for all NDPs including ADP, GDP, CDP and UDP (which converts to dUDP and in a different reaction to dTDP). The reaction is as follows: [thioredoxin]-dithiol + ribonucleoside 5'-diphosphate ↔ [thioredoxin]-disulfide + 2'-deoxyribonucleoside 5'-diphosphate + H2O To refresh your mind, thioredoxin is a small protein (12K) that is part of a complex with thioredoxin reductase and thioredoxin-interacting protein. It has two key sulfhydryls at the active site which act as reducing agents as they get converted to a disulfide as shown in Figure \(12\). Figure \(12\): Reduced thioredoxin and its oxidized form This single enzyme is so critical to cellular life that we will examine it more closely. There are several classes of these enzymes (Ia-Ie, II, and III). The class I enzymes generally use a di-transition metal complex as a cofactor while class II uses adenosylcobalamin. We will focus on class I enzymes, which have two subunits, called α and β or M1 and M2, respectively. The NDP binds in an α/M1 subunit active site which is developed only in the dimer. The β/M2 subunit is often referred to as the radical-generating subunit as it contains the transition metal complex that generates a free tyrosyl radical cation critical to the reaction. Almost all class I ribonucleotide reductases (RNRs) use transition metal ions located in the β/M2 subunit in the catalytic cycle for the dehydroxylation of the 2' OH on the ribose ring of the nucleotide. The metal ion complex a β/M2 subunit tyrosine to a tyrosine free radical cation which oxidizes an active site cysteine in the α/M1 subunit to form a thiol radical cation (Cys•+), called a thiyl radical. This abstracts a H• from the 3'C on the ribose of the substrate, forming a 3'C radical cation. This facilitates a dehydration reaction which leads to the dehydroxylation of the 2' OH, regenerating the thiyl radical. Reducing equivalents to restore the catalytic function of the enzyme come from the oxidation of a thioredoxin disulfide bound in the other subunit of the protein or a formate. Given the importance of these enzymes, they must be highly regulated. There are two regulatory sites: • a specificity site: determines nucleotide (NDP) specificity • an activity site: regulates catalytic activity The specificity and activity sites are in the α/M1 subunit where allosteric regulators dNTPs and ATP bind to different sites. When ATP is bound, the enzyme uses CDP and UDP as substrates. When dGTP is bound, ADP is the preferred substrate. Finally, when dTTP is bound, GDP is the preferred substrate. The enzyme is inhibited by dATP binding to the actual active site. Figure \(13\)s shows an abbreviated mechanism and cartoon showing the activities of the two subunits. Figure \(13\): Abbreviated mechanism and a cartoon showing the activities of the two subunits. in class I RNR. A, universal mechanism for nucleotide reduction in RNRs. B, diagram of the steps involved in radical translocation in class I RNRs. Ruskoski and. Boal. J. Biol. Chem. (2021) 297(4) 101137. DOI:https://doi.org/10.1016/j.jbc.2021.101137. CC-BY license (http://creativecommons.org/licenses/by/4.0/) Radical formation starts at tyrosine 122 in the metal center site in the β/M2 subunit. Electron transfer then occurs across the two subunits from a very distant active site Cys 439 in the α subunit. which enables the formation of the thiyl radical cation (Cys•+). The structure of an E. Coli Type IA enzyme with bound ligands has been determined after much effort that involved trapping of a long-life intermediate. It required replacement of Tyr 122 in both β chains with 2,3,5-trifluorotyrosine, which allowed the structure to be determined by cyro-EM. Tyr 122 in the β chain forms starts the process of electron transfer as it becomes the Tyr 122.+ radical cation. A detailed mechanisms showing both electron transfer to Y122.+ and accompanying proton transfer is shown in Figure \(14\). Figure \(14\): Pathway of electron and proton transfers for the formation of the thiyl radical cation (Cys•+). after Kang et al. Science (2020). DOI: 10.1126/science.aba6794) The path for electron flow from the sulfur of C439 in the β subuit to regenerate Y122 is shown. That electron transfer occurs over a very long distance of 35 Å as it hops from the original sulfur donor to the acceptor. Figure \(15\) shows an interactive iCn3D model of the holocomplex of E. coli class Ia ribonucleotide reductase with GDP and TTP (6W4X). Figure \(15\): Holocomplex of E. coli class Ia ribonucleotide reductase with GDP and TTP (6W4X). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...71ubUDRovXURG8 The structure is a tetramer of two α chains (different shades of gray) and two β chains (different shades of cyan). The GDP and TTP in the α subunit and the Fe cluster in the β subunit are shown in CPK spacefill and labeled. The key side chains in the α and β chains that participate in electron transfer over the 35 Å distance are shown in CPK-colored sticks and labeled. The Fe2+ ions are in the form of a μ-oxo-diron complex (2 Fe ions coordinated by 1 oxide). Figure \(16\) shows specificity and catalytic sites of the biologically active tetramer form for Class 1 RNR as well as the transition metal cofactors (Fe, Mn, or both) in various Class 1 RNRs. Figure \(16\): Quaternary structure of the active holoenzyme complex in class I RNR (PDB accession code 6W4X). Insets show the location of the active site in the catalytic α subunit (middle top) and the metallo- or radical cofactor (middle bottom and far right) in the β subunit The tyrosyl free radical forms on binding of dioxygen to the transition metal ion center electron and subsequent loss of an electron from tyrosine, as shown in Figure \(17\): Figure \(17\): Cofactor assembly mechanisms for class I RNRs. Manganese-dependent enzymes are highlighted in purple. Superoxide-dependent RNRs are highlighted in hot pink. Subclasses that require a NrdI activase are indicated with a yellow box. Metal-centered Cys oxidants shown in green and Tyr-derived radical Cys oxidants shown in blue. The mechanisms of cofactor actions are not fully understood. O2 initially adds to the Fe2+/Fe2+ cluster which forms a peroxo-Fe3+/Fe3+ intermediate. Structural features of the active site for different class I RNRs along with redox-active transition metal ions are shown in Figure \(18\). Figure \(18\): Comparison of metal-binding sites in class I RNRs The PDB codes for each structure are as follows: A, (3N3A), B, 4M1I), C, (6CWP), and D (6EBO). Water molecules are shown as red spheres. Now let's look at the regulation of class IA RNRs in more detail. Let's consider perhaps the most important allosteric regulators which bind 15 Å from the active site which affect RNR enzymatic activity: • dATP: inhibits RNRs when it binds to the α subunit • ATP: reverses the inhibition by dATP dATP/ATP also affects RNR enzyme specificity as they tilt the preference of RNR towards pyrimidine substrates, where TTP and dGTP promote purine substrate binding. These same rules apply to human and E. coli RNRs, with the locations of the active sites also being the same. The allosteric regulators appear to affect the quaternary structure of the enzyme. In E. Coli, the binding of dADP converts the structure of the enzyme from an active α2β2 form to an inhibited α4β4 ring structure. When dATP is bound, a "cone" domain in α forms interactions with the β subunit, leading to the formation of a dimer of the α2β2 tetramer. ATP reverses this effect by displacing dATP and pushing the equilibrium towards the active α2β2 form. This is shown in Panel A of Figure \(19\). Figure \(19\): Comparison of mechanisms of allosteric regulation of activity for E. coli and human RNRs. Brignole et al. 3.3-Å resolution cryo-EM structure of human ribonucleotide reductase with substrate and allosteric regulators bound eLife 7:e31502. https://doi.org/10.7554/eLife.31502. Attribution 4.0 International (CC BY 4.0). Panel B above illustrates the regulation of the human enzyme, which appears to be quite different. In the absence of either dATP or ATP, RNRs exist just as α2 dimers. On binding either dATP or ATP, the α2 dimer is converted to an α6 (a trimer of dimers) structure. Both are inactive in the absence of β subunits (which provide the metal cofactor site) • When β2 is added to the dATP bound α6 hexamer, the hexamer becomes stabilized, but the resulting complex is inhibited. • When β2 is added to the ATP bound α6 hexamer, the hexamer becomes destabilized and into smaller structures which are active. Hence the ratio of cellular dATP/ATP changes the aggregation state of the RNR and hence its activity in both E. Coli and humans. Figure \(20\) shows an interactive iCn3D model of the Human ribonucleotide reductase large subunit (alpha) with dATP and CDP (6AUI). Figure \(20\): Human ribonucleotide reductase large subunit (alpha) with dATP and CDP (6AUI). (Copyright; author via source). Click the image for a popup or use this external link:https://structure.ncbi.nlm.nih.gov/i...w1sFwwYSAD4nG6 Each of the α subunits in the hexamer is shown in a different color. dATPs (allosteric inhibitors) are shown in spacefill, red. CDPs (substrates) are shown in spacefill yellow. The hole in the middle of the structure prevents β2 from binding in a catalytically productive fashion, so the structure is inactive. A specific loop, loop 2, assists in the determination of RNRs specificity. It is between the base of dATP and the base of the substrate CDP. The backbone of the loop interacts with the adenine base and orients Gln 288 in the active site to interact with the cytosine of the substrate CDP. In both systems, backbone atoms of this loop ‘read out’ the adenine base and position Gln288 into the active site to recognize the cytosine of CDP. Figure \(20\) gives further details on the origin of substrate specificity. Figure \(21\): Determinants of substrate specificity are conserved from E. coli to humans. Panel (A) shows residues of human α (blue) interacting with CDP (carbons in orange) in the active site and dATP (carbons in yellow) in the specificity site. Density for CDP in orange mesh and for dATP in the yellow mesh. Panel (B) zooms in on dATP in the specificity site. Water molecules and oxygen atoms are in red, nitrogen in blue, magnesium in green, and phosphate in gold. Panel (C) zooms in on CDP in the active site. Panel (D) shows an overlay of human α from the α6 EM structure in blue with E. coli α from the α4β4-CDP-dATP cocrystal structure in gray (PDB: 5CNS) shows a nearly identical loop 2 conformation positioning Gln288 and Arg293 (Gln294 and Arg298 in E. coli). Panel (E) shows an overlay of human α from the α6 EM structure in blue with the crystal structure of human α with N- and C-termini truncated (residues 77–742) cocrystallized with dATP in tan (PDB: 2WGH) shows similar positioning of dATP but an altered conformation of loop 2 in the absence of bound CDP. The CDP shown is from the α6 EM structure. Panel (F) shows an overlay of human α from the α6 EM structure in blue with equivalent residues of yeast α structure with CDP and AMPPNP in brown (PDB: 2CVU) and shows a conformation of loop 2 that is distinct from that seen in structures of E. coli and human α Transcriptional regulation of RNRs Given the importance of this key enzyme, it should come as no surprise that its levels are regulated at the transcription level. As the activity of RNR is determined by its polymeric quaternary state, the transcriptional activation of the genes for RNRs is controlled in bacteria by quaternary states of the RNR-specific transcriptional repressor NrdR. The transcription factors bind to a specific DNA sequence called an NrdR box, which precedes the start site of transcription for RNR, where RNA polymerase binds. The NrdR protein acts to repress the synthesis of RNR genes. Its aggregation state is controlled by dATP/ATP ratios. When dATP is high, the NrdR binds to DNA and represses the synthesis of the RNR gene. In contrast when ATP is high, the protein does not bind to DNA and hence it can not repress transcription. The association /dissociation of the repressor is controlled by the aggregation state of NrdR. When abundant, NrdR exists as a 12-mer complex with two molecules of ATP bound per monomer. This ATP-bound 12-mer can't bind DNA, so transcription of the gene for RNR is not repressed. As dATP increases, one ATP is displaced so each monomer has 1 dATP and 1 ATP bound. This causes the NrdR to covet to an 8-mer. A 4-mer (tetrameric) version of this protein binds to the NrdR box sequence at the start of the RNR gene, repressing its synthesis. Figure \(22\) shows the dodecameric, octameric, and tetrameric structures of NrdR and their functions. Figure \(22\): Mechanism of NrdR function involving dodecameric, octameric, and tetrameric structures. Rozman Grinberg, I., Martínez-Carranza, M., Bimai, O. et al. A nucleotide-sensing oligomerization mechanism that controls NrdR-dependent transcription of ribonucleotide reductases. Nat Commun 13, 2700 (2022). https://doi.org/10.1038/s41467-022-30328-1. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/. Panel (a) shows a surface representation of the cryo-EM maps for the dodecameric, octameric, and DNA-bound tetrameric NrdR structures. Panel (b) shows a cartoon representation of the ATP-loaded NrdR tetramer (left) and the dATP/ATP-loaded tetramer (right). Chains A, B, C, and D are colored in beige, green, pink, and blue, respectively. Panel (c) shows the interface between the ATP cones in chain A (beige) and chain B (green) in the ATP-loaded dodecamer. Panel (d) shows the dATP/ATP-loaded tetramer. Panels c, and d were made from the same perspective, based on an alignment of the ATP-cones in chains A and B in both structures. Figure \(23\) shows an interactive iCn3D model of the Streptomyces coelicolor dATP/ATP-loaded NrdR in complex with its cognate DNA (7P3F). Figure \(23\): Streptomyces coelicolor dATP/ATP-loaded NrdR in complex with its cognate DNA (7P3F). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...2u3HMRPmJHe9X9 The monomers of the NrdR tetramer are shown in different colors. The yellow spheres are dATP and the orange ones are ATP. Pyrimidine Salvage Pathway There is also a salvage pathway as shown in Figure \(24\). Figure \(24\): Pyrimidine salvage pathway (after Wang et al. Frontiers in Oncology, 11 (2021). https://www.frontiersin.org/article/...nc.2021.684961. DOI=10.3389/fonc.2021.684961 Cells at rest use the salvage pathway reactants derived from nucleic acid degradation generic nucleoside pools. Nucleotide Degradation Purines The pathway for purine degradation is shown in Figure \(25\). Figure \(25\): Purine degradation Pyrimidine Degradation The pathway for pyrimidine degradation is shown in Figure \(26\). Figure \(26\): Pyrimidine degradation

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/02%3A_Unit_II-_Bioenergetics_and_Metabolism/22%3A_Biosynthesis_of_Amino_Acids_Nucleotides_and_Related_Molecules/22.04%3A_Biosynthesis_and_Degradation_of_Nucle.txt

• 23.1: Gene Mapping and Chromosomal Karyotypes Genes provide instructions to build living organisms and each specific gene maps to the same chromosome in every cell. This physical gene location within the organism's chromosomes is called the gene loci. If two genes are found on the same chromosome, especially when they are in close proximity to one another, they are said to be linked. • 23.2: DNA Transposable Elements Eukaryotic genomes contain an abundance of repeated DNA, and some repeated sequences are mobile. Transposable elements (TEs) are defined as DNA sequences that are able to move from one location to another in the genome. TEs have been identified in all organisms, prokaryotic and eukaryotic, and can occupy a high proportion of a species’ genome. • 23.3: Chromosome Packaging Most eukaryotic chromosomes include packaging proteins which, aided by chaperone proteins, bind to and condense the DNA molecule to prevent it from becoming an unmanageable tangle. Before typical cell division, these chromosomes are duplicated in the process of DNA replication, providing a complete set of chromosomes for each daughter cell. 23: Chromosome Structure Search Fundamentals of Biochemistry Introduction Genes provide instructions to build living organisms and each specific gene maps to the same chromosome in every cell. This physical gene location within the organism's chromosomes is called the gene loci. If two genes are found on the same chromosome, especially when they are near one another, they are said to be linked. Genetic linkage is the tendency of DNA sequences that are close together on a chromosome to be inherited together during the meiosis phase of sexual reproduction. Two genetic markers that are physically near to each other are unlikely to be separated into different chromatids during chromosomal crossover and are therefore said to be more linked than markers that are far apart. In other words, the nearer two genes are on a chromosome, the lower the chance of recombination between them, and the more likely they are to be inherited together. Markers on different chromosomes are perfectly unlinked. Genetic linkage is the most prominent exception to Gregor Mendel's Law of Independent Assortment. The first experiment to demonstrate linkage was carried out in 1905. At the time, the reason why certain traits tend to be inherited together was unknown. Later work revealed that genes are physical structures related by physical distance. The typical unit of genetic linkage is the centimorgan (cM). A distance of 1 cM between two markers means that the markers are separated into different gametes on average once per 100 meiotic products, thus once per 50 meioses. A linkage map (also known as a genetic map) is a table for a species or experimental population that shows the position of its known genes or genetic markers relative to each other in terms of recombination frequency, rather than a specific physical distance along each chromosome. Linkage maps were first developed by Alfred Sturtevant, a student of Thomas Hunt Morgan. Figure \(1\) shows a gene linkage map with the relative positions of allelic characteristics on the second Drosophila chromosome. A linkage map is a map based on the frequencies of recombination between markers during the crossover of homologous chromosomes. The greater the frequency of recombination (segregation) between two genetic markers, the further apart they are assumed to be. Conversely, the lower the frequency of recombination between the markers, the smaller the physical distance between them. Historically, the markers originally used were detectable phenotypes (enzyme production, eye color) derived from coding DNA sequences; eventually, confirmed or assumed noncoding DNA sequences such as microsatellites or those generating restriction fragment length polymorphisms (RFLPs) have been used. Linkage maps help researchers to locate other markers, such as other genes by testing for genetic linkage of the already known markers. In the early stages of developing a linkage map, the data are used to assemble linkage groups, a set of genes that are known to be linked. As knowledge advances, more markers can be added to a group, until the group covers an entire chromosome. For well-studied organisms, the linkage groups correspond one-to-one with the chromosomes. Traditional studies used to physically map genes onto specific chromosomes were painstaking and involved using restriction enzymes to fragment the genome of an organism and then clone the fragments into YACs or BACs creating a DNA library. The library could then be screened with specific genetic probes to determine which fragment contained a gene of interest. The fragments would then need to be sequenced and reassembled using overlapping patterns. Today, the sequencing of entire genomes from nearly any organism is possible and relatively easy in comparison. Thus, a traditional genetic map can more readily be overlayed on the physical chromosomal map of an organism as shown in Figure \(2\). This was one of the overarching goals of the human genome project. Karyotypes The entire chromosome set of a species is known as a karyotype, which can be thought of as a global map of the nuclear genome. Karyotyping is the process by which the condensed chromosomes of an organism are stained and photographed using light microscopy. Karyotyping can be used to determine the chromosome complement of an individual, including the number of chromosomes and any abnormalities. Karyotypes describe the chromosome count of an organism and what these chromosomes look like under a light microscope. Attention is paid to their length, the position of the centromeres, banding pattern, any differences between the sex chromosomes, and any other physical characteristics. The preparation and study of karyotypes are part of the larger field of cytogenetics. The field of cytogenetics involves the study of inheritance in relation to the structure and function of chromosomes. Thus, karyotyping is a fundamental process within this field. The study of whole sets of chromosomes is sometimes known as karyology. The chromosomes are depicted (by rearranging a photomicrograph) in a standard format known as a karyogram or idiogram: in pairs, ordered by size and position of centromere for chromosomes of the same size, as shown in Figure \(3\). The basic number of chromosomes in the somatic cells of an individual or a species is called the somatic number and is designated 2n. In the germ line (the sex cells) the chromosome number is n (humans: n = 23). Thus, in humans 2n = 46. In normal diploid organisms, autosomal chromosomes are present in two copies. There may, or may not, be sex chromosomes. Polyploid cells have multiple copies of chromosomes and haploid cells, usually, gametes, have single copies. Karyotypes can be used for many purposes; such as to study chromosomal aberrations, cellular function, taxonomic relationships, medicine and to gather information about past evolutionary events (karyosystematics). During the chromosomal staining processes used to produce a karyotype, the staining intensity along the chromosome can vary due to localized sequence and structural differences. These banding patterns are an inherent characteristic of a chromosome and can be utilized as a diagnostic tool. Typically, karyotypes are prepared from cells that are actively undergoing mitosis. The mitotic progression is blocked in prometaphase or metaphase when chromosomes exist in their most condensed state. The cells are lysed, but the nuclei are retained intact and are subsequently treated with a chemical fixing agent. Once fixed, a number of different types of stains can be used to visualize the chromosomes. One of the first types of chromosomal staining procedures developed was known as Q-banding, which was developed in 1970 by Torbjorn Caspersson. This technique uses the DNA-alkylating dye, quinacrine, which forms a covalent link with the DNA. Researchers noted that the staining patterns resulting from this technique were consistent and repeatable, demonstrating that banding patterns can be used to identify and characterize individual chromosomes. Giemsa dye, as shown in Figure 24.3, is more commonly used today, as it can be used with bright field microscopy and produces high detail banding patterns. A specific technique, called G-banding uses Giemsa staining following the treatment of mitotic chromosomes with the protease, trypsin. Pre-treating the sample with trypsin before staining causes the partial breakdown of chromosomal proteins leading to chromosomal relaxation. This allows more thorough staining of the chromosomes when treated with Giemsa dye. When the chromosomal region is more tightly packed into heterochromatin, it tends to stain more darkly with the Giemsa dye, than the more lightly packaged euchromatic regions. Heterochromatic regions tend to have higher A-T content and don't contain as many gene regions as euchromatic regions. Euchromatic regions stain more lightly with G-banding. Other types of staining with Giemsa, include R-banding or Reverse-banding, which involves heating the DNA before staining. This is thought to cause the melting of A-T-rich regions, reducing the Giemsa staining, when compared to G-C-rich, gene-containing regions of the chromosomes. When visualizing a karyotype, the chromosomal images are aligned so that heterologous chromosomes are paired together and positioned such that the p-arm (short arm) is on top and the q-arm (long arm) points downward. Karyotypes can be used to quickly identify gross chromatic abnormalities that are larger than a few megabases in difference. This includes abnormalities such as aneuploidy (the addition or absence of an entire chromosome), or translocations (the transfer of part of a chromosome to a neighboring chromosome), as shown in Figure \(4\) and Figure \(5\). The telomeric regions of chromosomes can also be identified using fluorescent staining techniques, as shown in Figure \(6\). The structure of telomeric chromosomal regions is described in section 24.3. More recently, techniques such as chromosome painting, use fluorescently labeled probes to hybridize with specific chromosomes or even specific gene regions of a chromosome. Karyotypes originating from this technique are called spectral karyotypes. This technique can be especially useful in identifying translocations that have occurred in human cells, as shown in Figure \(7\).

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/23%3A_Chromosome_Structure/23.01%3A_Gene_Mapping_and_Chromosomal_Karyotypes.txt

Search Fundamentals of Biochemistry Introduction Eukaryotic genomes contain an abundance of repeated DNA, and some repeated sequences are mobile. Transposable elements (TEs) are defined as DNA sequences that can move from one location to another in the genome. TEs have been identified in all organisms, prokaryotic and eukaryotic, and can occupy a high proportion of a species’ genome. For example, transposable elements comprise approximately 10% of several fish species, 12 % of the C. elegans genome, 37% of the mouse genome, 45% of the human genome, and up to >80% of the genome of some plants like maize. From bacteria to humans, transposable elements have accumulated over time and continue to shape genomes through their mobilization. TEs were discovered by Barbara McClintock during experiments conducted in 1944 on maize. Since they appeared to influence phenotypic traits, she named them controlling elements. However, her discovery was met with less than enthusiastic reception by the genetic community. Her presentation at the 1951 Cold Spring Harbor Symposium was not understood and at least not very well received. She had no better luck with her follow-up publications and after several years of frustration decided not to publish on the subject for the next two decades. Not for the first time in the history of science, an unappreciated discovery was brought back to life after some other discovery has been made. In this case, it was the discovery of insertion sequences (IS) in bacteria by Szybalski group in the early 1970s. In the original paper, they wrote: “Genetic elements were found in higher organisms which appear to be readily transposed from one to another site in the genome. Such elements, identifiable by their controlling functions, were described by McClintock in maize. It is possible that they might be somehow analogous to the presently studied IS insertions”. The importance of McClintock’s original work was eventually appreciated by the genetic community with numerous awards, including 14 honorary doctoral degrees and a Nobel Prize in 1983 “for her discovery of mobile genetic elements”. Her picture is shown in Figure \(1\). The mobilization of TEs is termed transposition or retrotransposition, depending on the nature of the intermediate used for mobilization. There are several ways in which the activity of TEs can positively and negatively impact a genome; for example, TE mobilization can promote gene inactivation, modulate gene expression or induce illegitimate recombination. Thus, TEs have played a significant role in genome evolution. For example, DNA transposons can inactivate or alter the expression of genes by insertion within introns, exons, or regulatory regions. In addition, TEs can participate in the reorganization of a genome by the mobilization of non-transposon DNA or by acting as recombination substrates. This recombination would occur by homology between two sequences of a transposon located in the same or different chromosomes, which could be the origin of several types of chromosome alterations. Indeed, TEs can participate in the loss of genomic DNA by internal deletions or other mechanisms. The reduction in fitness suffered by the host due to transposition ultimately affects the transposon, since host survival is critical to the perpetuation of the transposon. Therefore, strategies have been developed by host and transposable elements to minimize the deleterious impact of transposition, and to reach equilibrium. For example, some transposons tend to insert in nonessential regions in the genome, such as heterochromatic regions, where insertions will likely have a minimal deleterious impact. In addition, they might be active in the germ line or embryonic stage, where most deleterious mutations can be selected against during fecundation or development, allowing only non-deleterious or mildly deleterious insertions to pass to successive generations. New insertions may also occur within an existing genomic insertion to generate an inactive transposon or can undergo self-regulation by overproduction-inhibition. On the other hand, host organisms have developed different mechanisms of defense against high rates of transposon activity, including DNA-methylation to reduce TE expression, several RNA interference-mediated mechanisms, mainly in the germ line, or through the inactivation of transposon activity by the action of specific proteins. In some cases, transposable elements have been “domesticated” by the host to perform a specific function in the cell. A well-known example is RAG proteins, which participate in V(D)J recombination during antibody class switching, and exhibit a high similarity to DNA transposons, from which these proteins appear to be derived. Another example is the centromeric protein CENP-B, which seems to have originated from the pogo-like transposon. The analogous human mariner Himar1 element has been incorporated into the SETMAR gene, which consists of the histone H3 methylase gene and the Himar1 transposase domain. This gene is involved in the non-homologous end-joining pathway of DNA repair and has been shown to confer resistance to ionizing radiation. From a genome-wide view, it has been estimated that ~25% of human promoter regions and ~4% of human exons contain sequences derived from TEs. Thus, we are likely underestimating the rate of domestication events in mammalian genomes. The first TE classification system was proposed by Finnegan in 1989 and distinguished two classes of TEs characterized by their transposition intermediate: RNA (class I or retrotransposons) or DNA (class II or DNA transposons). The transposition mechanism of class I is commonly called “copy and paste” and that of class II, “cut and paste.” In 2007 Wicker et al. proposed a hierarchical classification based on TEs structural characteristics and mode of replication, as shown in Figure \(2\). Class I: Mobile Elements As mentioned above, class I TEs transpose through an RNA intermediary. The RNA intermediate is transcribed from genomic DNA and then reverse-transcribed into DNA by a TE-encoded reverse transcriptase (RT), followed by reintegration into a genome. Each replication cycle produces one new copy, and as a result, class I elements are the major contributors to the repetitive fraction in large genomes. Retrotransposons are divided into five orders: LTR retrotransposons, DIRS-like elements, Penelope-like elements (PLEs), LINEs (long interspersed elements), and SINEs (short interspersed elements). This scheme is based on the mechanistic features, organization, and reverse transcriptase phylogeny of these retroelements. Accidentally, the retrotranscriptase coded by an autonomous TE can reverse-transcribe another RNA present in the cell, e.g., mRNA, and produce a retrocopy of it, which in most cases results in a pseudogene. The LTR retrotransposons are characterized by the presence of long terminal repeats (LTRs) ranging from several hundred to several thousand base pairs. Both exogenous retroviruses and LTR retrotransposons contain a gag gene that encodes a viral particle coat and a pol gene that encodes a reverse transcriptase, ribonuclease H, and an integrase, which provide the enzymatic machinery for reverse transcription and integration into the host genome. Reverse transcription occurs within the viral or viral-like particle (GAG) in the cytoplasm, and it is a multi-step process. Unlike LTR retrotransposons, exogenous retroviruses contain an env gene, which encodes an envelope that facilitates their migration to other cells. Some LTR retrotransposons may contain remnants of an env gene, but their insertion capabilities are limited to the originating genome. This would rather suggest that they originated in exogenous retroviruses by losing the env gene. However, there is evidence that suggests the contrary, given that LTR retrotransposons can acquire the env gene and become infectious entities. Presently, most of the LTR sequences (85%) in the human genome are found only as isolated LTRs, with the internal sequence being lost most likely due to homologous recombination between flanking LTRs. Interestingly, LTR retrotransposons target their reinsertion to specific genomic sites, often around genes, with putative important functional implications for a host gene. It is estimated that 450,000 LTR copies make up about 8% of our genome. LTR retrotransposons inhabiting large genomes, such as maize, wheat, or barley, can contain thousands of families. However, despite the diversity, very few families comprise most of the repetitive fraction in these large genomes. Notable examples are Angela (wheat), BARE1 (barley), Opie (maize), and Retrosor6 (sorghum). The DIRS order clusters structurally diverged groups of transposons that possess a tyrosine recombinase (YR) gene instead of an integrase (INT) and do not form target site duplications (TSDs). Their termini resemble either split direct repeats (SDR) or inverted repeats. Such features indicate a different integration mechanism than that of other class I mobile elements. DIRS were discovered in the slime mold (Dictyostelium discoideum) genome in the early 1980s, and they are resent in all major phylogenetic lineages including vertebrates. It has been shown that they are also common in hydrothermal vent organisms. Another order, termed Penelope-like elements (PLE), has wide, though patchy distribution from amoebae and fungi to vertebrates with copy numbers up to thousands per genome. Interestingly, no PLE sequences have been found in mammalian genomes, and apparently, they were lost from the genome of C. elegans. Although PLEs with an intact ORF have been found in several genomes, including Ciona and Danio, the only transcriptionally active representative, Penelope, is known from Drosophila virilis. It causes the hybrid dysgenesis syndrome characterized by the simultaneous mobilization of several unrelated TE families in the progeny of dysgenic crosses. It seems that Penelope invaded D. virilis quite recently, and its invasive potential was demonstrated in D. melanogaster. PLEs harbor a single ORF that codes for a protein containing reverse transcriptase (RT) and endonuclease (EN) domains. The PLE RT domain more closely resembles telomerase than the RT from LTRs or LINEs. The EN domain is related to GIY-YIG intron-encoded endonucleases. Some PLE members also have LTR-like sequences, which can be in a direct or an inverse orientation, and have a functional intron. LINEs do not have LTRs; however, they have a poly-A tail at the 3′ ends and are flanked by the TSDs. They comprise about 21% of the human genome and among them L1 with about 850,000 copies is the most abundant and best-described LINE family. L1 is the only LINE retroposon still active in the human genome. In the human genome, there are two other LINE-like repeats, L2 and L3, distantly related to L1. A contrasting situation has been noticed in the malaria mosquito Anopheles gambiae, where around 100 divergent LINE families compose only 3% of its genome. LINEs in plants, e.g., Cin4 in maize and Ta11 in Arabidopsis thaliana, seem rare as compared with LTR retrotransposons. A full copy of mammalian L1 is about 6 kb long and contains a PolII promoter and two ORFs. The ORF1 codes for a non-sequence-specific RNA binding protein that contains zinc finger, leucine zipper, and coiled-coil motifs. The ORF1p functions as a chaperone for the L1 mRNA. The second ORF encodes an endonuclease, which makes a single-stranded nick in the genomic DNA, and a reverse transcriptase, which uses the nicked DNA to prime reverse transcription of LINE RNA from the 3′ end. Reverse transcription is often unfinished, leaving behind fragmented copies of LINE elements; hence most of the L1-derived repeats are short, with an average size of 900 bp. LINEs are part of the CR1 clade, which has members in various metazoan species, including fruit flies, mosquito, zebrafish, pufferfish, turtles, and chicken. Because they encode their own retrotransposition machinery, LINE elements are regarded as autonomous retrotransposons. SINEs evolved from RNA genes, such as 7SL and tRNA genes. By definition, they are short, up to 1000 base pairs long. They do not encode their own retrotranscription machinery and are considered nonautonomous elements and in most cases are mobilized by the L1 machinery. The outstanding member of this class from the human genome is the Alu repeat, which contains a cleavage site for the AluI restriction enzyme that gave its name. With over a million copies in the human genome, Alu is probably the most successful transposon in the history of life. Primate-specific Alu and its rodent relative B1 have limited phylogenetic distribution suggesting their relatively recent origins. The mammalian-wide interspersed repeats (MIRs), by contrast, spread before eutherian radiation, and their copies can be found in different mammalian groups including marsupials and monotremes. SVA elements are unique primate elements due to their composite structure. They are named after their main components: SINE, VNTR (a variable number of tandem repeats), and Alu. Usually, they contain the hallmarks of the retroposition, i.e., they are flanked by TSDs and terminated by a poly(A) tail. It seems that SVA elements are nonautonomous retrotransposons mobilized by L1 machinery, and they are thought to be transcribed by RNA polymerase II. SVAs are transpositionally active and are responsible for some human diseases. They originated less than 25 million years ago, and they form the youngest retrotransposon family with about 3000 copies in the human genome. Retro(pseudo)genes are a special group of retroposed sequences, which are products of reverse transcription of a spliced (mature) mRNA. Hence, their characteristic features are an absence of promoter sequence and introns, the presence of flanking direct repeats, and a 3′-end polyadenosine tract. Processed pseudogenes, as sometimes retropseudogenes are called, have been generated in vitro at a low frequency in the human HeLa cells via mRNA from a reporter gene. The source of the reverse transcription machinery in humans and other vertebrates seems to be active L1 elements. However, not all retroposed messages have to end up as pseudogenes. About 20% of mammalian protein-encoding genes lack introns in their ORFs. It is conceivable that many genes lacking introns arose by retroposition. Some genes are known to be retroposed more often than others. For instance, in the human genome, there are over 2000 retropseudogenes of ribosomal proteins. A genome-wide study showed that the human genome harbors about 20,000 pseudogenes, 72% of which most likely arose through retroposition. Interestingly, the vast majority (92%) of them are quite recent transpositions that occurred after primate/rodent divergence. Some of the retroposed genes may undergo quite complicated evolutionary paths. An example could be the RNF13B retrogene, which replaced its own parental gene in the mammalian genomes. This retrocopy was duplicated in primates, and the evolution of this primate-specific copy was accompanied by the exaptation of two TEs, Alu and L1, and intron gain via changing a part of the coding sequence into an intron leading to the origin of a functional, primate-specific retrogene with two splicing variants. Class II: Mobile Elements Class II elements move by a conservative cut-and-paste mechanism; the excision of the donor element is followed by its reinsertion elsewhere in the genome. DNA transposons are abundant in bacteria, where they are called insertion sequences, but are also present in all phyla. Two subclasses of DNA transposons have been distinguished, based on the number of DNA strands that are cut during transposition. Classical “cut-and-paste” transposons belong to subclass I, and they are classified as the TIR order. They are characterized by terminal inverted repeats (TIR) and encode a transposase that binds near the inverted repeats and mediates mobility. This process is not usually a replicative one, unless the gap caused by excision is repaired using the sister chromatid. When inserted at a new location, the transposon is flanked by small gaps, which, when filled by host enzymes, cause duplication of the sequence at the target site. The length of these TSDs is characteristic of particular transposons. Nine superfamilies belong to the TIR order, including Tc1-Mariner, Merlin, Mutator, and PiggyBac. The second-order Crypton consists of a single superfamily of the same name. Originally thought to be limited to fungi, now it is clear that they have a wide distribution, including animals and heterokonts. A heterogeneous, small, nonautonomous group of elements MITEs also belong to the TIR order, which in some genomes amplified to thousands of copies, e.g., Stowaway in the rice genome, Tourist in most bamboo genomes, or Galluhop in the chicken genome. Subclass II includes two orders of TEs that, just as those from subclass I, do not form RNA intermediates. However, unlike “classical” DNA transposons, they replicate without double-strand cleavage. Helitrons replicate using a rolling-circle mechanism, and their insertion does not result in the target site duplication. They encode tyrosine recombinase along with some other proteins. Helitrons were first described in plants, but they are also present in other phyla, including fungi and mammals. Mavericks are large transposons that have been found in different eukaryotic lineages excluding plants. They encode various numbers of proteins that include DNA polymerase B and an integrase. Kapitonov and Jurka suggested that their life cycle includes a single-strand excision, followed by extrachromosomal replication and reintegration to a new location. TEs are not randomly distributed in the genome As seen in the previous section, TEs are highly diverse and in principle, every TE sequence in a genome can be affiliated to a (sub)family, superfamily, subclass, and class. This is summarized in Figure \(3\). However, much like the taxonomy of species, the classification of TEs is in constant flux, perpetually subject to revision due to the discovery of completely novel TE types, the introduction of new levels of granularity in the classification, and the ongoing development of methods and criteria to detect and classify TEs. The genome may be viewed as an ecosystem inhabited by diverse communities of TEs, which seek to propagate and multiply through sophisticated interactions with each other and with other components of the cell. These interactions encompass processes familiar to ecologists, such as parasitism, cooperation, and competition. Thus, it is perhaps not surprising that TEs are rarely, if ever, randomly distributed in the genome, as shown in Figure \(4\). TEs exhibit various levels of preference for insertion within certain features or compartments of the genome. These are often guided by opposite selective forces, a balancing act of facilitating future propagation while mitigating deleterious effects on host cell function. At the most extreme end of the site-selection spectrum, many elements have evolved mechanisms to target specific loci where their insertions are less detrimental to the host but favorable for their propagation. For instance, several retrotransposons in species as diverse as slime mold and budding and fission yeast have evolved independently, but convergently, the capacity to target the upstream regions of genes transcribed by RNA polymerase III, where they do not appear to affect host gene expression but retain the ability to be transcribed themselves. TEs are an extensive source of mutations and genetic polymorphisms TEs occupy a substantial portion of the genome of a species, including a large fraction of the DNA unique to that species. In maize, where Barbara McClintock did her seminal work, an astonishing 60 to 70% of the genome is comprised of LTR retrotransposons, many of which are unique to this species or its close wild relatives, but the less prevalent DNA transposons are currently the most active and mutagenic (Fig. 24.2.4). Similarly, the vast majority of TE insertions in Drosophila melanogaster are absent at the orthologous site in its closest relative D. simulans (and vice versa), and most are not fixed in the population. Many TE families are still actively transposing and the process is highly mutagenic; more than half of all known phenotypic mutants of D. melanogaster isolated in the laboratory are caused by spontaneous insertions of a wide variety of TEs. Transposition events are also common and mutagenic in laboratory mice, where the ongoing activity of several families of LTR elements are responsible for 10–15% of all inherited mutant phenotypes. This contribution of TEs to genetic diversity may be underestimated, as TEs can be more active when organisms are under stress, such as in their natural environment. Because TE insertions rarely provide an immediate fitness advantage to their host, those reaching fixation in the population do so largely by genetic drift and are subsequently eroded by point mutations that accumulate neutrally. Over time, these mutations result in TEs that can no longer encode transposition enzymes and produce new integration events. For instance, our (haploid) genome contains ~ 500,000 L1 copies, but more than 99.9% of these L1 copies are fixed and no longer mobile due to various forms of mutations and truncations. It is estimated that each person carries a set of ~ 100 active L1 elements, and most of these are young insertions still segregating within the human population. Thus, as for any other organism, the ‘reference’ human genome sequence does not represent a comprehensive inventory of TEs in humans. Thousands of ‘non-reference’, unfixed TE insertions have been cataloged through whole genome sequencing and other targeted approaches. On average, any two human haploid genomes differ by approximately a thousand TE insertions, primarily from the L1 or Alu families. The number of TE insertion polymorphisms in a species with much higher TE activity such as maize dwarfs the number in humans. If TEs bring no immediate benefit to their host and are largely decaying neutrally once inserted, how do they persist in evolution? One key to this conundrum is the ability of TEs not only to propagate vertically but also horizontally between individuals and species. There is now a large body of evidence supporting the idea that horizontal transposon transfer is a common phenomenon that affects virtually every major type of TE and all branches of the tree of life. While the cellular mechanisms underlying horizontal transposon transfer remain murky, it is increasingly apparent that the intrinsic mobility of TEs and ecological interactions between their host species, including those with pathogens and parasites, facilitate the transmission of elements between widely diverged taxa. TEs are associated with genome rearrangements and unique chromosome feature Transposition represents a potent mechanism of genome expansion that over time is counteracted by the removal of DNA via deletion. The balance between the two processes is a major driver in the evolution of genome size in eukaryotes. Several studies have demonstrated the impact and range of this shuffling and cycling of genomic content on the evolution of plant and animal genomes. Because the insertion and removal of TEs are often imprecise, these processes can indirectly affect surrounding host sequences. Some of these events occur at high enough frequency to result in vast amounts of duplication and reshuffling of host sequences, including genes and regulatory sequences. For example, a single group of DNA transposons (MULEs) has been responsible for the capture and reshuffling of ~ 1,000 gene fragments in the rice genome. Such studies have led to the conclusion that the rate at which TEs transpose, which is in part under host control, is an important driver of genome evolution. In addition to rearrangements induced as a byproduct of transposition, TEs can promote genomic structural variation long after they have lost the capacity to mobilize. In particular, recombination events can occur between the highly homologous regions dispersed by related TEs at distant genomic positions and result in large-scale deletions, duplications, and inversions (Fig. 24.2.4). TEs also provide regions of microhomology that predispose to template switching during repair of replication errors leading to another source of structural variants. These non-transposition-based mechanisms for TE-induced or TE-enabled structural variation have contributed substantially to genome evolution. These processes can also make the identification of actively transposing elements more difficult in population studies that infer the existence of active elements through the detection of non-reference insertions. TEs also contribute to specialized chromosome features. An intriguing example is in Drosophila, where LINE-like retrotransposons form and maintain the telomeres in replacement of the telomerase enzyme which has been lost during dipteran evolution. This domestication event could be viewed as a replay of what might have happened much earlier in eukaryotic evolution to solve the ‘end problem’ created by the linearization of chromosomes. Indeed, the reverse transcriptase component of telomerase is thought to have originated from an ancient lineage of retroelements. TE sequences and domesticated transposase genes also play structural roles at centromeres. There is an intricate balance between TE expression and repression To persist in evolution, TEs must strike a delicate balance between expression and repression (Fig. 24.2.4). Expression should be sufficient to promote amplification, but not so vigorous as to lead to a fitness disadvantage for the host that would offset the benefit to the TE of increased copy numbers. This balancing act may explain why TE-encoded enzymes are naturally suboptimal for transposition and why some TEs have evolved self-regulatory mechanisms controlling their own copy numbers. A variety of host factors are also employed to control TE expression, which includes a variety of small RNA, chromatin, and DNA modification pathways, as well as sequence-specific repressors such as the recently profiled KRAB zinc-finger proteins. However, many of these silencing mechanisms must be at least partially released to permit the developmental regulation of host gene expression programs, particularly during early embryonic development. For example, genome-wide loss of DNA methylation is necessary to reset imprinted genes in primordial germ cells. This affords TEs an opportunity, as reduced DNA methylation often promotes TE expression. Robust expression of a TE in the germ lineage (but not necessarily in the gametes themselves) is often its own downfall. In one example of a clever trick employed by the host, TE repression is relieved in a companion cell derived from the same meiotic product as flowering plant sperm. However, this companion cell does not contribute genetic material to the next generation. Thus, although TEs transpose in a meiotic product, the events are not inherited. Instead, TE activity in the companion cell may further dampen TE activity in sperm via the import of TE-derived small RNAs. Another important consequence of the intrinsic expression/repression balance is that the effects of TEs on a host can vary considerably among tissue types and stages of an organism’s life cycle. From the TE’s perspective, an ideal scenario is to be expressed and active in the germline, but not in the soma, where expression would gain the TE no advantage, only disadvantages. This is indeed observed among many species, with ciliates representing an extreme example of this division—TEs are actively deleted from the somatic macronucleus but retained in the micronucleus, or germline. Another example is the P-elements in Drosophila, which are differentially spliced in the germline versus soma. Many organisms, including plants, do not differentiate germ lineage cells early in development; rather, they are specified from somatic cells shortly before meiosis commences. Thus, TEs that transpose in somatic cells in plants have the potential to be inherited, which suggests that the interest of TEs and hosts are in conflict across many more cells and tissues than in animals with a segregated germline. TEs are insertional mutagens in both germline and soma Like other species, humans contend with a contingent of currently active TEs where the intrinsic balance between expression and repression is still at play. For us, this includes L1 and other mobile elements that depend on L1-encoded proteins for retrotransposition. These elements are responsible for new germline insertions that can cause genetic disease. More than 120 independent TE insertions have been associated with human disease. The rate of de novo germline transposition in humans is approximately one in 21 births for Alu and one in 95 births for L1. Historically, little attention has been given to transposition in somatic cells and its consequences, because somatic transposition may be viewed as an evolutionary dead-end for the TE with no long-term consequences for the host species. Yet, there is abundant evidence that TEs are active in somatic cells in many organisms (Fig. 24.2.4). In humans, L1 expression and transposition have been detected in a variety of somatic contexts, including early embryos and certain stem cells. There is also a great deal of interest in mobile element expression and activity in the mammalian brain, where L1 transposition has been proposed to diversify neuronal cell populations. One challenge for assessing somatic activity has rested with the development of reliable single-cell insertion site mapping strategies. Somatic activity has also been observed in human cancers, where tumors can acquire hundreds of new L1 insertions. Just like for human polymorphisms, somatic activity in human cancers is caused by small numbers of so-called ‘hot’ L1 loci. The activities of these master copies vary depending on the individual, tumor type, and timeframe in the clonal evolution of the tumor. Some of these de novo L1 insertions disrupt critical tumor suppressors and oncogenes and thus drive cancer formation, although the vast majority appear to be ‘passenger’ mutations. Host cells have evolved several mechanisms to keep TEs in check. However, as the force of natural selection begins to diminish with age and completely drops in post-reproductive life, TEs may become more active. TEs can be damaging in ways that do not involve transposition TEs are best known for their mobility, and their ability to transpose to new locations. While the breakage and insertion of DNA associated with transposition represent an obvious source of cell damage, this is not the only or perhaps even the most common mechanism by which TEs can be harmful to their host. Reactivated transposons harm the host in multiple ways. First, de-repression of transposon loci, including their own transcription, may interfere with transcription or processing of host mRNAs through a myriad of mechanisms. Genome-wide transcriptional de-repression of TEs has been documented during replicative senescence of human cells and several mouse tissues, including the liver, muscle, and brain. De-repression of LTR and L1 promoters can also cause oncogene activation in cancer. Second, TE-encoded proteins such as the endonuclease activity of L1 ORF2p can induce DNA breaks and genomic instability. Third, accumulation of RNA transcripts and extrachromosomal DNA copies derived from TEs may trigger an innate immune response leading to autoimmune diseases and sterile inflammation (Fig. 24.2.4). Activation of interferon response is now a well-documented property of transcripts derived from endogenous retroviruses and may give immunotherapies a boost in identifying and attacking cancer cells. The relative contribution of all the above mechanisms in organismal pathologies remains to be determined. Following transcription (and sometimes splicing) of TEs, the next step in the process involves the translation of the encoded proteins and, for retroelements, reverse transcription of the TEs into cDNA substrates suitable for transposition. Once engaged by a TE-encoded reverse transcriptase protein, the resulting cytosolic DNAs and RNA:DNA hybrids can alert inflammatory pathways. An example of this is seen in patients with Aicardi–Goutières syndrome, where the accumulation of TE-derived cytosolic DNA is due to mutations in pathways that normally block TE processing or degrade TE-derived DNA. Although not all TEs encode functional proteins, some do, including a few endogenous retroviruses capable of producing Gag, Pol, or envelope (Env) proteins. Overexpression of these Env proteins can be cytotoxic and has been linked to at least two neurodegenerative diseases, multiple sclerosis, and amytrophic lateral sclerosis. Small accessory proteins produced by the youngest human endogenous retrovirus (HERV) group, HERV-K (HML-2), may play a role in some cancers but the evidence remains circumstantial. Key coding and non-coding RNAs are derived from TEs Although usually detrimental, there is growing evidence that TE insertions can provide the raw material for the emergence of protein-coding genes and non-coding RNAs, which can take on important and, in some cases essential, cellular function (Fig. 24.2.4). The process of TE gene ‘domestication’ or exaptation over evolutionary time contributes to both deeply conserved functions and more recent, species-specific traits. Most often, the ancestral or a somewhat modified role of a TE-encoded gene is harnessed by the host and conserved, while the rest of the TE sequence, and hence its ability to autonomously transpose, has been lost. Spectacular examples of deeply conserved TE-derived genes are Rag1 and Rag2, that catalyze V(D)J somatic recombination in the vertebrate immune system. Both genes, and probably the DNA signals they recognize, were derived from an ancestral DNA transposon around 500 million years ago. Indeed, DNA transposases have been co-opted multiple times to form new cellular genes. The gag and env genes of LTR retrotransposons or endogenous retroviruses (ERVs) have also been domesticated numerous times to perform functions in placental development, contribute to host defense against exogenous retroviruses, act in brain development, and play other diverse roles. One of the most intriguing examples of TE domestication is the repeated, independent capture of ERV env genes, termed syncytins, which appear to function in placentation by facilitating cell–cell fusion and syncytiotrophoblast formation. Notably, one or more syncytin genes have been found in virtually every placental mammalian lineage where they have been sought, strongly suggesting that ERVs have played essential roles in the evolution and extreme phenotypic variability of the mammalian placenta. Another example of a viral-like activity re-purposed for host cell function is provided by the neuronal Arc gene, which arose from the gag gene from a LTR retrotransposon domesticated in the common ancestor of tetrapod vertebrates. Genetic and biochemical studies of murine Arc show that it is involved in memory and synaptic plasticity and has preserved most of the ancestral activities of Gag, including the packaging and intercellular trafficking of its own RNA. Remarkably, flies appear to have independently evolved a similar system of trans-synaptic RNA delivery involving a gag-like protein derived from a similar yet distinct lineage of LTR retrotransposons. Thus, the biochemical activities of TE-derived proteins have been repeatedly co-opted during evolution to foster the emergence of convergent cellular innovations in different organisms. TEs can donate their own genes to the host, but they can also add exons and rearrange and duplicate existing host genes. In humans, intronic Alu elements are particularly prone to be captured as alternative exons through cryptic splice sites residing within their sequences. L1 and SVA (SINE/VNTR/Alu) elements also contribute to exon shuffling through transduction events of adjacent host sequences during their mobilization. The reverse transcriptase activity of retroelements is also responsible for the trans-duplication of cellular mRNAs to create ‘processed’ retrogenes in a wide range of organisms. The L1 enzymatic machinery is thought to be involved in the generation of tens of thousands of retrogene copies in mammalian genomes, many of which remain transcribed and some of which have acquired new cellular functions. This is a process still actively shaping our genomes; it has been estimated that 1 in every 6000 humans carries a novel retrogene insertion. TEs also make substantial contributions to the non-protein coding functions of the cell. They are major components of thousands of long non-coding RNAs in human and mouse genomes, often transcriptionally driven by retroviral LTRs. Some of these TE-driven lncRNAs appear to play important roles in the maintenance of stem cell pluripotency and other developmental processes. Many studies have demonstrated that TE sequences embedded within lncRNAs and mRNAs can directly modulate RNA stability, processing, or localization with important regulatory consequences. Furthermore, TE-derived microRNAs and other small RNAs processed from TEs can also adopt regulatory roles serving host cell functions. The myriad of mechanisms by which TEs contribute to coding and non-coding RNAs illustrate the multi-faceted interactions between these elements and their host. TEs contribute cis-regulatory DNA elements and modify transcriptional networks Cis-regulatory networks coordinate the transcription of multiple genes that function in concert to orchestrate entire pathways and complex biological processes. In line with Barbara McClintock’s insightful predictions, there is now mounting evidence that TEs have been a rich source of material for the modulation of eukaryotic gene expression (Fig. 24.2.4). Indeed, TEs can disperse vast amounts of promoters and enhancers, transcription factor binding sites, insulator sequences, and repressive elements. The varying coat colors of agouti mice provide a striking example of a host gene controlling coat color whose expression can be altered by the methylation levels of a TE upstream of its promoter. In the oil palm, the methylation level of a TE that sits within a gene important for flowering ultimately controls whether or not the plants bear oil-rich fruit. As TE families typically populate a genome as a multitude of related copies, it has long been postulated that they have the potential to donate the same cis-regulatory module to ‘wire’ batteries of genes dispersed throughout the genome. An increasing number of studies support this model and suggest that TEs have provided the building blocks for the assembly and remodeling of cis-regulatory networks during evolution, including pathways underlying processes as diverse as pregnancy, stem cell pluripotency, neocortex development, innate immunity in mammals, or the response to abiotic stress in maize. Indeed, TE sequences harbor all the necessary features of a ‘classical’ gene regulatory network. They are bound by diverse sets of transcription factors that integrate multiple inputs (activation/repression), respond to signals in both cis and trans, and are capable of co-ordinately regulating gene expression. In this context, TEs are highly suitable agents to modify biological processes by creating novel cis-regulatory circuits and fine-tuning pre-existing networks. Outlook As potent insertional mutagens, TEs can have both positive and negative effects on host fitness, but it is likely that the majority of TE copies in any given species—and especially those such as humans with small effective population size—have reached fixation through genetic drift alone and are now largely neutral to their host. When can we say that TEs have been co-opted for cellular function? The publication of the initial ENCODE paper, which asserted ‘function for 80% of the genome’, was the subject of much debate and controversy. Technically speaking, ENCODE assigned only ‘biochemical’ activity to this large fraction of the genome. Yet critics objected to the grand proclamations in the popular press (The Washington Post Headline: “Junk DNA concept debunked by new analysis of the human genome”) and to the ENCODE consortium’s failure to prevent this misinterpretation. To these critics, ignoring evolutionary definitions of function was a major misstep. This debate can be easily extended to include TEs. TEs make up the vast majority of what is often referred to as ‘junk DNA’. Today, the term is mostly used (and abused) by the media, but it has deep roots in evolutionary biology. Regardless of the semantics, what evidence is needed to assign a TE with a function? Many TEs encode a wide range of biochemical activities that normally benefit their own propagation. For example, TEs often contain promoter or enhancer elements that highjack cellular RNA polymerases for transcription and autonomous elements encode proteins with various biochemical and enzymatic activities, all of which are necessary for the transposon to replicate. Do these activities make them functional? The vast differences in TEs between species make standard approaches to establishing their regulatory roles particularly challenging. For example, intriguing studies on the impact of HERVs, in particular HERV-H, in stem cells and pluripotency must be interpreted using novel paradigms that do not invoke deep evolutionary conservation to imply function, as these particular ERVs are absent outside of great apes. Evolutionary constraints can be measured at shorter time scales, including the population level, but this remains a statistically challenging task, especially for non-coding sequences. Natural loss-of-function alleles may exist in the human population and their effect on fitness can be studied if their impact is apparent, but these are quite rare and do not allow systematic studies. It is possible to engineer genetic knockouts of a particular human TE locus to test its regulatory role but those are restricted to in-vitro systems, especially when the orthologous TE does not exist in the model species. In this context, studying the impact of TEs in model species with powerful genome engineering tools and vast collections of mutants and other genetic resources, such as plants, fungi, and insects, will also continue to be extremely valuable. Finally, a growing consensus is urging more rigor when assigning cellular function to TEs, particularly for the fitness benefit of the host. Indeed, a TE displaying biochemical activity (such as those bound by transcription factors or lying within open chromatin regions) cannot be equated to a TE that shows evidence of purifying selection at the sequence level or, when genetically-altered, result in a deleterious or dysfunctional phenotype. Recent advances in editing and manipulating the genome and the epigenome en masse yet with precision, including repetitive elements, offer the promise for a systematic assessment of the functional significance of TEs.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/23%3A_Chromosome_Structure/23.02%3A_DNA_Transposable_Elements.txt

Search Fundamentals of Biochemistry Some of the material in this chapter section comes from Chapter 8.4, Chromosomes and Chromatin, as it was important to describe it earlier in the structure/function unit. In addition, some biochemistry courses might not get to the material in a late chapter in a text. Repetition of some of the material is easier in an online textbook as well. Introduction Recall from Chapter 8, that within eukaryotic cells, DNA is organized into long linear structures called chromosomes, as shown in Figure \(1\). A chromosome is a deoxyribonucleic acid (DNA) molecule with part or all of the genetic material (genome) of an organism. Most eukaryotic chromosomes include packaging proteins which, aided by chaperone proteins, bind to and condense the DNA molecule to prevent it from becoming an unmanageable tangle. Before typical cell division, these chromosomes are duplicated in the process of DNA replication, providing a complete set of chromosomes for each daughter cell. The replicated arms of a chromosome are called chromatids. Before being separated into the daughter cells during mitosis, replicated chromatids are held together by a chromosomal structure called the centromere. Eukaryotic organisms (animals, plants, fungi, and protists) store most of their DNA inside the cell nucleus as linear nuclear DNA, and some in the mitochondria as circular mitochondrial DNA or in chloroplasts as circular chloroplast DNA. In contrast, prokaryotes (bacteria and archaea) do not have organelle structures and thus, store their DNA only in a region of the cytoplasm known as the nucleoid region. Prokaryotic chromosomes consist of double–stranded circular DNA. The genome of a cell is often significantly larger than the cell itself. For example, if the DNA from a human cell containing 46 chromosomes were stretched out in a line, it would extend more than 6 feet (2 meters)! How is it possible that the genetic information not only fits into the cell but fits into the cell nucleus? Eukaryota solves this problem by a combination of supercoiling and packaging DNA around the histone family of proteins (described below). Prokaryotes do not contain histones (with a few exceptions). Prokaryotes tend to compress their DNA using nucleoid-associated-proteins (NAPs) and supercoiling (Figure 24.3.2). Supercoiling DNA supercoiling refers to the over- or under-winding of a DNA strand, and is an expression of the strain on that strand, as shown in Figure \(2\). Supercoiling is important in many biological processes, such as compacting DNA, and regulating access to the genetic code. DNA supercoiling strongly affects DNA metabolism and possibly gene expression. Additionally, certain enzymes such as topoisomerases can change DNA topology to facilitate functions such as DNA replication or transcription. In a “relaxed” double-helical segment of B-DNA, the two strands twist around the helical axis once every 10.4–10.5 base pairs of sequence. Adding or subtracting twists, as some enzymes can do, impose strain. If a DNA segment under twist strain were closed into a circle by joining its two ends and then allowed to move freely, the circular DNA would contort into a new shape, such as a simple figure-eight (Figure \(2\)). Such a contortion is a supercoil. The noun form “supercoil” is often used in the context of DNA topology. Positively supercoiled (overwound) DNA is transiently generated during DNA replication and transcription, and, if not promptly relaxed, inhibits (regulates) these processes. The simple figure eight is the simplest supercoil and is the shape a circular DNA assumes to accommodate one too many or one too few helical twists. The two lobes of the figure- eight will appear rotated either clockwise or counterclockwise with respect to one another, depending on whether the helix is over- or underwound. For each additional helical twist being accommodated, the lobes will show one more rotation about their axis. As a general rule, the DNA of most organisms is negatively supercoiled. Lobal contortions of a circular DNA, such as the rotation of the figure-eight lobes above, are referred to as writhe. The above example illustrates that twist and writhes are interconvertible. Supercoiling can be represented mathematically by the sum of twist and writhe (Figure \(2\). The twist is the number of helical turns in the DNA and the writhe is the number of times the double helix crosses over on itself (these are the supercoils). Extra helical twists are positive and lead to positive supercoiling, while subtractive twisting causes negative supercoiling. Many topoisomerase enzymes sense supercoiling and either generate or dissipate it as they change DNA topology. In addition to forming supercoiled structures, circular chromosomes from bacteria have been shown to undergo the processes of catenation and knotting upon the inhibition of topoisomerase enzymes. Catenation is the process by which two circular DNA strands are linked together like chain links, whereas DNA knotting is the interlooping structures occurring within a single circular DNA structure, as shown in Figure \(3\). In vivo, the action of topoisomerase enzymes is critical to keep knots and catenoids from tangling the DNA structure. Catenanes are effectively topologically linked circular molecules In part, because chromosomes may be very large, segments in the middle may act as if their ends are anchored. As a result, they may be unable to distribute excess twist to the rest of the chromosome or to absorb twist to recover from underwinding—the segments may become supercoiled, in other words. In response to supercoiling, they will assume an amount of writhe, just as if their ends were joined. Supercoiled circular DNA forms two major structures; a plectoneme or a toroid, or a combination of both (Figure 24.3.2). A negatively supercoiled DNA molecule will produce either a one-start left-handed helix, the toroid, or a two-start right-handed helix with terminal loops, the plectoneme. Plectonemes are typically more common in nature, and this is the shape most bacterial plasmids will take (Figure 4.10). For larger molecules, it is common for hybrid structures to form – a loop on a toroid can extend into a plectoneme, as shown in Figure \(4\). DNA supercoiling is important for DNA packaging within all cells, and seems to also play a role in gene expression. Topoisomerases Topoisomerase can change the tension in supercoiled DNA. Think of how you untie a knot. It takes a lot of work sometimes, and if it's too hard, you simply cut the impediment to unknotting. Topoisomerases work by making transient breaks in the DNA before unwinding and religation. There are two main types of topoisomerases, topo I and topo II. It's very hard to describe their activities with just words and static diagrams. View the video below and you will get a great sense of what the enzymes do and how they are different. With this background, we can now explore each enzyme in more detail. They are both targets of cancer drugs which makes them even more interesting. Topo I enzyme relaxes DNA by nicking one stand. The dsDNA then rotates around the non-nicked strand. It unwinds new DNA and allows the condensation of chromosomes. When both DNA and RNA polymerase makes new DNA and RNA strands, respectively, they increase the supercoiling of the nucleic acid. Topoisomerases relax them. They also play a role in the regulation of gene expression by affecting gene promoters where RNA polymerase binds, with negative supercoiling enhancing transcription and positive supercoiling inhibiting it. Figure \(5\) shows the topology of DNA and an overview of the mechanisms of Topo I and II. : DNA topology and DNA topoisomerase mechanism Figure \(5\): DNA topology and DNA topoisomerase mechanisms. Shannon J. McKie, Keir C. Neuman, and Anthony Maxwell. Bioessays (2021). https://doi.org/10.1002/bies.202000286. Attribution 4.0 International (CC BY 4.0) (A) Topological consequences of DNA metabolism. i) During DNA replication, strand separation leads to positive supercoiling ahead of the advancing protein machinery, and precatenane formation behind. Precatenanes form as the newly-synthesized duplexes wrap around one another, and, if not removed before completion of replication, catenated DNA molecules are formed. ii) During transcription, strand separation leads to positive supercoiling ahead of the advancing protein machinery, and negative supercoil formation behind. iii) Hemicatenanes are a possible end result of replication, in which the parental strands of the replicated duplexes remain base-paired. iv: DNA knotting can also occur as a result of DNA replication in which a DNA molecule is intramolecularly linked. (B) Summary of topo categories and mechanism. The topos are categorized based on whether they catalyze single- (type I) or double-stranded (type II) DNA breaks. The type I topos are further subdivided to type IA, IB, and IC. Type IA form a transient covalent bond to the 5ʹ DNA phosphate and function via a strand passage mechanism. Type IB and IC form a transient covalent bond to the 3ʹ DNA phosphate and function via a controlled-rotation mechanism. Type II topos are further subdivided into type IIA and IIB. Both form a transient covalent bond to the 5ʹ DNA phosphate of both strands of the duplex and function via a strand-passage mechanism. (C) Summary of the topological manipulations performed by DNA topoisomerases, namely relaxation of positive and negative supercoils and decatenation. Type IA topos are color-coded pink, type IB are orange, type IC are yellow, type IIA are green, and type IIB are blue. The requirement of ATP or ssDNA for activity is denoted using a red or blue circle, respectively Topoisomerase I (Topo I): Class I topoisomerases wrap around the DNA and cut one strand. Keeping that spot in place, the helix can spin to reduce strain caused by either over- or underwinding. After these geometric contortions, the single-stranded DNA nick is repaired and the tension is relieved. Type IA topoisomerase from E. Coli is shown in Figure \(6\). Figure \(6\): Structure of a type IA topoisomerase. E. coli topoisomerase III is shown to illustrate the overall structure of a type IA topoisomerase and the typical toroidal fold observed in all members of this type. (A) Diagram showing the structure of the apo-enzyme [PDB 1D6M(7)]. In the absence of DNA, the active site, found at the intersection of domains I and III (encircled), is buried. (B) Diagram showing the structure of a complex with single-stranded DNA [PDB 1I7D (23)]. Note the movement of domains that occurs to accommodate DNA. In both diagrams, the four major domains of the protein are colored red, blue, purple, and green for domains I, II, III, and IV, respectively. The single-stranded DNA binding groove, shown circled in black, extends from domain IV to the active site. The active site residues as well as the single-stranded DNA in the complex are shown in a ball and stick representation. Dasgupta T, Ferdous S, Tse-Dinh YC. Mechanism of Type IA Topoisomerases. Molecules. 2020 Oct 17;25(20):4769. doi: 10.3390/molecules25204769. PMID: 33080770; PMCID: PMC7587558. Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/). A simplified cartoon mechanism for Topo I is shown in Figure \(7\). Figure \(7\): Diagram showing the proposed mechanism of DNA relaxation by type IA topoisomerases. The mechanism involves several transient conformational intermediates both of the protein and the DNA. The sequence of the steps and the intermediates are hypothetical and more states are likely to be involved in the cycle. Processivity by the enzyme requires that after one relaxation event, the protein continues to another relaxation cycle without releasing the DNA. In the diagram, the protein is shown in grey, and the DNA in red/blue. The orange dot represents the presence of the covalent protein/DNA complex. The single-stranded DNA binding groove is shown in red or yellow. Dasgupta, T et al. ibid. A more detailed view of the domain structure and mechanism for Topo I is shown in Figure \(8\). Figure \(8\): Type IA DNA topoisomerases. Dasgupta, T et al. ibid. (A) Protein domain organization of Escherichia coli DNA topoisomerase IA (topo IA) and DNA topoisomerase III (topo III). Black vertical lines represent the active site tyrosines. (B) Crystal structure of E. coli topo I bound to ssDNA (PDB: 4RUL).[20] (C) Strand-passage mechanism for type IA topos. (1) topo binds the G-segment ssDNA region, (2) the G-segment is cleaved. (3) The topo DNA gate is opened, (4) which allows T-segment transfer through the cleaved G-strand. (5) The DNA gate is closed, (6) and the G-strand is re-ligated, changing the linking number by 1. (7) The topo can then go through another round of relaxation or dissociate from the DNA. Type IA topo (domains 1–4) is in pink, the active site tyrosine is yellow and the DNA is grey. (D) Crystal structure of E. coli topo III bound to ssDNA (PDB: 2O54).[26] (E) Crystal structures of human topo IIIα (blue) bound to RMI1(orange) (PDB: 4CGY),[39] and human topo IIIβ (magenta) bound to TDRD3 (green) (PDB: 5GVE).[60] For panels A, B, and C, the topo I and III domains are color-coded as follows: D1 is red, D2 is pink, D3 is yellow, D4 is orange, D5 is marine blue, D6 is purple, D7 is green, D8 is teal, and D9 is light blue Figure \(9\) shows an interactive iCn3D model of E.Coli topoisomerase I in complex with ssDNA (4RUL). Figure \(9\): E.Coli topoisomerase I in complex with ssDNA (4RUL). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...JxcFvds2e8p2WA The topoisomerase is shown in gray and the single-stranded DNA is in cyan. 4 Zn2+ ions are shown. Three form Zn-finger motifs. Two of these are shown near the protein binding site for DNA. A fourth Zn is bound to a single His 566 in the structure. The active site amino acids of the enzyme, which includes residues D111, D113, Y319, and R321 from D1 and D3, are shown and labeled. Also shown are active site residues, E115, F139 and Y312. There are 4 Cys-Zn ribbon domains. The ones that interact with the ssDNA involve π-stacking, some of which are illustrated in the iCn3D model. Class II topoisomerases (Topo II) A series of enzymes are included in this class including DNA gyrase and Topo (IV) from prokaryotes and Topo II from eukaryotes. In eukaryotes, they help sister chromosomes separate if they get tangled during cell division. This enzyme works by: this enzyme makes a double-stranded cut, moves on the helix through the cut, and reseals the cut. • binding the gate segment (G -segment) ds-DNA at a DNA gate where a double-stranded break is made • binding the transport segment (T-segment) ds-DNA at the N-gate where the nucleotide ATP binds • The T-segment DNA moves through the break in the G-segment and released the C-gate • The G- and T-segments are reconnected. After this, the N gate reopens to allow the process to occur again. The domain structure of Topo IIs and the general mechanism of action are shown in Figure \(10\). Figure \(10\): Type II DNA topoisomerases: domain organization and mechanism. (A) Protein domain organization for the type IIA topos: E. coli DNA gyrase, E. coli DNA topoisomerase IV (topo IV), yeast DNA topoisomerase II (topo II), Methanosarcina mazei DNA topoisomerase VI (topo VI), Paenibacillus polymyxa DNA topoisomerase VIII (plasmid-borne), and Pseudomonas phage NP1 Mini-A. (B) type II topo strand passage mechanism. (1) G-segment is bound at the DNA gate and the T-segment is captured. (2) ATP binding stimulates dimerization of the N-gate, the G-segment is cleaved and the T-segment is passed through the break. (3) The G-segment is re-ligated and T-segment exits through the C-gate. For type IIB topos, there is no C-gate so once the T-segment passes through the G-segment, it is released from the enzyme. (4) Dissociation of ADP and Pi allows N-gate opening, a scenario where the enzyme either remains bound to the G-segment, ready to capture a consecutive T-segment, or (5) dissociates from the G-segment. Figure \(11\) shows an interactive iCn3D model of Yeast Topoisomerase II-DNA-AMPPNP complex (4GFH) . Figure \(11\): Yeast Topoisomerase II-DNA-AMPPNP complex (4GFH). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...b5EHLfMwde5Dw8 The two protein chains in the homodimer are shown in magenta and cyan. The double-stranded DNA is shown with a brown backbone and CPK-colored spheres. ANP (AMPPNP), a nonhydrolyzable ATP analog, and Mg2+ ions are shown in spacefill and labeled. The ATP binding and ATPase domain of one of the monomers (cyan for example) is adjacent to the nuclease-cutting domain of the other monomer (magenta). This requires some conformational gymnastics as the ATP binding and cleavage domain move around each other to allow the DNA strand to pass in the right direction and to reset the enzyme. Note the circular nature of chloroplast and mitochondrial DNA, suggesting a bacterial origin for both of these organelle structures. Sequence alignments further lend support for the endosymbiotic theory, which proposes that bacteria were engulfed by early eukaryotic organisms and subsequently became symbiotic to their eukaryotic counterpart, rather than being digested. A reminder about mitochondrial DNA In the cells of extant organisms, the vast majority of the proteins present in the mitochondria (numbering approximately 1500 different types in mammals) are coded for by nuclear DNA. However, sequencing of the human mitochondrial genome has revealed 16,569 base pairs encoding 13 proteins (Figure 24.3.5). Many of the mitochondrially produced proteins are required for electron transport during the production of ATP, as shown in Figure \(12\). Histones and DNA packing Within eukaryotic chromosomes, chromatin proteins, known as histones, compact and organize DNA. These compacting structures guide the interactions between DNA and other proteins, helping control which parts of the DNA are transcribed. Histones are highly alkaline proteins found in eukaryotic cell nuclei that package and order the DNA into structural units called nucleosomes. They are the chief protein components of chromatin, acting as spools around which DNA winds, and playing a role in gene regulation. Without histones, the unwound DNA in chromosomes would be very long (a length-to-width ratio of more than 10 million to 1 in human DNA). For example, each human diploid cell (containing 23 pairs of chromosomes) has about 1.8 meters of DNA; wound on the histones, the diploid cell has about 90 micrometers (0.09 mm) of chromatin. Five major families of histones exist: H1/H5, H2A, H2B, H3, and H4. Histones H2A, H2B, H3, and H4 are known as the core histones, while histones H1/H5 are known as the linker histones. The core histones all exist as dimers, which are similar in that they all possess the histone fold domain: three alpha helices linked by two loops. It is this helical structure that allows for interaction between distinct dimers, particularly in a head-tail fashion (also called the handshake motif). The resulting four distinct dimers then come together to form one octameric nucleosome core, approximately 63 Angstroms in diameter. Around 146 base pairs (bp) of DNA wrap around this core particle 1.65 times in a left-handed super-helical turn to give a particle of around 100 Angstroms across, called a nucleosome, as shown in Figure \(13\). The linker histone H1 binds the nucleosome at the entry and exit sites of the DNA, thus locking the DNA into place and allowing the formation of a higher order structure, as shown in Figure \(14\). The most basic such formation is the 10 nm fiber or beads on a string conformation. This involves the wrapping of DNA around nucleosomes with approximately 50 base pairs of DNA separating each pair of nucleosomes (also referred to as linker DNA). The nucleosome contains over 120 direct protein-DNA interactions and several hundred water-mediated ones. Direct protein – DNA interactions are not spread evenly about the octamer surface but rather located at discrete sites. These are due to the formation of two types of DNA binding sites within the octamer; the α1α1 site, which uses the α1 helix from two adjacent histones, and the L1L2 site formed by the L1 and L2 loops. Salt links and hydrogen bonding between both side-chain basic and hydroxyl groups and main-chain amides with the DNA backbone phosphates form the bulk of interactions with the DNA. This is important, given that the ubiquitous distribution of nucleosomes along genomes requires it to be a non-sequence-specific DNA-binding factor. Although nucleosomes tend to prefer some DNA sequences over others, they are capable of binding practically to any sequence, which is thought to be due to the flexibility in the formation of these water-mediated interactions. In addition, non-polar interactions are made between protein side-chains and the deoxyribose groups, and an arginine side-chain intercalates into the DNA minor groove at all 14 sites where it faces the octamer surface. The distribution and strength of DNA-binding sites about the octamer surface distort the DNA within the nucleosome core. The DNA is non-uniformly bent and also contains twist defects. The twist of free B-form DNA in solution is 10.5 bp per turn. However, the overall twist of nucleosomal DNA is only 10.2 bp per turn, varying from a value of 9.4 to 10.9 bp per turn. The histone tail extensions constitute up to 30% by mass of the histones but are not visible in the crystal structures of nucleosomes due to their high intrinsic flexibility, and have been thought to be largely unstructured (Figure 4.14). The N-terminal tails of histones H3 and H2B pass through a channel formed by the minor grooves of the two DNA strands, protruding from the DNA every 20 bp. The N-terminal tail of histone H4, on the other hand, has a region of highly basic amino acids (16-25), which, in the crystal structure, forms an interaction with the highly acidic surface region of a H2A-H2B dimer of another nucleosome, being potentially relevant for the higher-order structure of nucleosomes. This interaction is thought to occur under physiological conditions also, and suggests that acetylation of the H4 tail distorts the higher-order structure of chromatin. Figure \(15\) shows an interactive iCn3D model of the human nucleosome (3afa). One member of each pair of histones is shown in cartoon rendering, while the other member of the pair is shown in the same color but in spacefill rendering. The structure of a human nucleosome (3afa) is shown below (H2A is shown in cyan, H2B in blue, H3 in magenta, and H4 in purple). Each strand of DNA is shown in a different shade of gray. Figure \(15\): Human nucleosome (3afa). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...B2SwQHYDLj4BJ6 The formation of the DNA double helix represents the first-order packaging of the chromosome structure. The formation of nucleosomes represents the second level of packaging for eukaryotic chromosomes. In vitro data suggests that nucleosomes are then arranged into either a solenoid structure which consists of 6 nucleosomes linked together by the Histone H1 linker proteins or a zigzag structure that is similar to the solenoid construct, as shown in Figure \(16\). Both the solenoid and zigzag structures are approximately 30 nm in diameter. The solenoid and zigzag structures reported from in vitro data have not yet been confirmed to occur in vivo. During interphase, each chromosome occupies a spatially limited, roughly elliptical domain which is known as a chromosome territory (CT). Each chromosome territory is comprised of higher-order chromatin units of ~1 Mb each. These units are likely built up from smaller loop domains that contain the solenoid/zigzag structural motifs. On the other hand, 1Mb domains can themselves serve as smaller units in higher-order chromatin structures. Chromosome territories are known to be arranged radially around the nucleus. This arrangement is both cell and tissue-type specific and is also evolutionarily conserved. The radial organization of chromosome territories was shown to correlate with their gene density and size. In this case, the gene-rich chromosomes occupy interior positions, whereas larger, gene-poor chromosomes, tend to be located around the periphery. Chromosome territories are also dynamic structures, with genes able to relocate from the periphery towards the interior once they have been ‘switched on’. In other cases, genes may move in the opposite direction, or simply maintain their position. The eviction of genes from their chromosome territories into the interchromatin compartment or a neighboring chromosome territory is often accompanied by the formation of large decondensed chromatin loops. Models describing chromosome territory arrangement With the development of high-throughput biochemical techniques, such as 3C (chromosome conformation capture) and 4C (chromosome conformation capture-on-chip and circular chromosome conformation capture), numerous spatial interactions between neighboring chromatin territories have been described, as shown in Figure \(17\). These descriptions have been supplemented with the construction of spatial proximity maps for the entire genome (e.g., for a human lymphoblastoid cell line). Together, these observations and physical simulations have led to the proposal of various models that aim to define the structural organization of chromosome territories: 1. The chromosome territory-interchromatin compartment (CT-IC) model describes two principal compartments: chromosome territories (CTs) and an interchromatin compartment (IC). In this model, chromosome territories build up an interconnected chromatin network that is associated with an adjacent 3D space called the interchromatin compartment. The latter can be observed using both light and electron microscopy. Within a single chromosome territory, the interphase chromosome is divided into defined regions based on the level of chromosome condensation. Here, the inner part of the interphase chromosome is comprised of more condensed chromatin domains or higher-order chromatin fibers, while a thin (<200 nm) layer of more decondensed chromatin, known as the perichromatin region, can be found around the chromosomal periphery. Functionally, the perichromatin region represents the major transcriptional compartment and is also the region where most co-transcriptional RNA splicing takes place. DNA replication and DNA repair are also predominately carried out within the perichromatin region. Finally, nascent RNA transcripts, referred to as perichromatin fibrils, are also generated in the perichromatin region. Perichromatin fibrils are then subjected to the splicing events by the factors, provided by the interchromatin compartment. The lattice model, proposed by Dehgani et al. is based on reports that transcription also occurs within the inner, more condensed chromosome territories and not only at the interface between the interchromatin compartment and the perichromatin region. Using ESI (electron spectroscopic imaging), Dehgani et al. showed that chromatin was organized as an array of deoxyribonucleoprotein fibers of 10–30 nm in diameter. In this study, the interchromatin compartments, which are described in the CT-IC model as large channels between chromosome territories, were not apparent. Instead, chromatin fibers created a loose meshwork of chromatin throughout the nucleus that intermingled at the periphery of chromosome territories. Thus, inter- and intra-chromosomal spaces within this meshwork are essentially contiguous and together form the intra-nuclear space. 2. The interchromatin network (ICN) model predicts that intermingling chromatin fibers/loops can make both cis- (within the same chromosome) and trans- (between different chromosomes) contacts. This intermingling is uniform and makes a distinction between the chromosome territory and interchromatin compartment functionally meaningless. The advantage of the ICN model is that it permits high chromatin dynamics and diffusion-like movements. The authors propose that ongoing transcription influences the degree of intermingling between specific chromosomes by stabilizing associations between particular loci. Such interactions are likely to depend on the transcriptional activity of the loci and are therefore cell-type specific. The cell type-specific organization of chromosome territories has been studied by measuring the volume and frequency of intermingling between heterologous chromosomes. By using 3C (chromosome conformation capture) and FISH (fluorescence in situ hybridization) to map the regions of chromosome intermingling, it was revealed that these regions contain a higher density of active genes and are enriched with markers of transcriptional activation and repression, such as activated RNAPII. By comparing the positions of the CTs in undifferentiated mouse embryonic stem (ES) cells, ES cells in early stages of differentiation, and terminally differentiated NIH3T3 cells, it was shown that fully differentiated cells had a higher enrichment of RNAPII, compared to undifferentiated or less-differentiated cells. The findings support the notion that the intermingling regions have functional significance in the nucleus and provide a basis for understanding how the radial and relative positions of chromosomal territories evolve during the process of differentiation, explaining their organization in a cell type-dependent manner. 3. The Fraser and Bickmore model emphasizes the functional importance of giant chromatin loops, which originate from chromosome territories and expand across the nuclear space to share transcription factories. In this case, both cis- and trans- oops of decondensed chromatin can be co-expressed and co-regulated by the same transcription factory. 4. The Chromatin polymer models assume a broad range of chromatin loop sizes and predict the observed distances between genomic loci and chromosome territories, as well as the probabilities of contacts being formed between given loci. These models apply physics-based approaches that highlight the importance of entropy for understanding nuclear organization. By proposing the existence of conformational chromatin ensembles with structures based on three possible homopolymer states, these models also provide alternative structures to the traditional 30 nm chromatin fiber, which has been brought into question following recent studies. With a lack of experimental evidence to support these described models, it must be remembered that they serve only to hypothesize the structural and chemical properties of intermediate chromatin structures and to highlight unanswered questions. For example, the mechanisms that exist to control the rate and the extent of chromatin movement remain to be defined At the ends of the linear chromosomes are specialized regions of DNA called telomeres, shown in Figure \(18\). The main function of these regions is to allow the cell to replicate chromosome ends using the enzyme telomerase, as the enzymes that normally replicate DNA cannot copy the extreme 3′ ends of chromosomes. These specialized chromosome caps also help protect the DNA ends, and stop the DNA repair systems in the cell from treating them as damage to be corrected. In human cells, telomeres are usually lengths of single-stranded DNA containing several thousand repeats of a simple TTAGGG sequence. In human cells, telomeres contain 300-8000 repeats of a simple TTAGGG sequence. The repetitive TTAGGG sequences in telomeric DNA can form unique higher-order structures called quadruplexes. Figure \(19\) shows an interactive iCn3D model of parallel quadruplexes from human telomeric DNA (1KF1). The structure contains a single DNA strand (5'-AGGGTTAGGGTTAGGGTTAGGG-3') which contains four TTAGGG repeats. Figure \(19\): A buried phenylalanine in low molecular weight protein tyrosyl phosphatase (1xww) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...y5joFHDgWJQsQ6 Rotate the model to see 3 parallel layers of quadruplexes. In each layer, 4 noncontiguous guanine bases interact with a K+ ion. Hover over the guanine bases in one layer and you will find that one layer consists of guanines 4, 10, 16, and 22, which derive from the last G in each of the repeats in the sequence of the oligomer used (5'-AGGGTTAGGGTTAGGGTTAGGG-3'). These quadruplexes certainly serve as recognition and the binding site for telomerase proteins. The guanine-rich telomere sequences which can form quadruplex may also function to stabilize chromosome ends During DNA replication, the double-stranded DNA is unwound and DNA polymerase synthesizes new strands. However, as DNA polymerase moves in a unidirectional manner (from 5’ to 3’), only the leading strand can be replicated continuously. In the case of the lagging strand, DNA replication is discontinuous. In humans, small RNA primers attach to the lagging strand DNA, and the DNA is synthesized in small stretches of about 100-200 nucleotides, which are termed Okazaki fragments. The RNA primers are removed, and replaced with DNA and the Okazaki fragments are ligated together. At the end of the lagging strand, it is impossible to attach an RNA primer, meaning that there will be a small amount of DNA lost each time the cell divides. This ‘end replication problem’ has serious consequences for the cell as it means the DNA sequence cannot be replicated correctly, with the loss of genetic information. To prevent this, telomeres are repeated hundreds to thousands of times at the end of the chromosomes. Each time cell division occurs, a small section of telomeric sequences is lost to the end replication problem, thereby protecting the genetic information. At some point, the telomeres become critically short. This attrition leads to cell senescence, where the cell is unable to divide, or apoptotic cell death. Telomeres are the basis for the Hayflick limit, the number of times a cell can divide before reaching senescence. Telomeres can be restored by the enzyme telomerase, which extends telomeres length (Figure 24.3.10). Telomerase activity is found in cells that undergo regular division, such as stem cells and lymphocyte cells of the immune system. Telomeres can also be extended through the Alternative Lengthening of Telomeres (ALT) pathway. In this case, rather than being extended, telomeres are switched between chromosomes by homologous recombination. As a result of the telomere swap, one set of daughter cells will have shorter telomeres, and the other set will have longer telomeres. A downside to telomere extension is the potential for uncontrolled cell division and cancer. Abnormally high telomerase activity has been found in the majority of cancer cells, and non-telomerase tumors often exhibit ALT pathway activation. As well as the potential for losing genetic information, cells with short telomeres are at high risk for improper chromosome recombination, which can lead to genetic instability and aneuploidy (an abnormal number of chromosomes). These guanine-rich telomere sequences may also stabilize chromosome ends by forming structures of stacked sets of four-base units, rather than the usual base pairs found in other DNA molecules (Figure 24.3.10). Here, four guanine bases form a flat plate and these flat four-base units then stack on top of each other, to form a stable G-quadruplex structure. These structures are stabilized by hydrogen bonding between the edges of the bases and the chelation of a metal ion in the center of each four-base unit. Other structures can also be formed, with the central set of four bases coming from either a single strand folded around the bases or several different parallel strands, each contributing one base to the central structure. In addition to these stacked structures, telomeres also form large loop structures called telomere loops or T-loops. Here, the single-stranded DNA curls around in a long circle stabilized by telomere-binding proteins. At the very end of the T-loop, the single-stranded telomere DNA is held onto a region of double-stranded DNA by the telomere strand disrupting the double-helical DNA and base pairing to one of the two strands. This triple-stranded structure is called a displacement loop or D-loop.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/23%3A_Chromosome_Structure/23.03%3A_Chromosome_Packaging.txt

Search Fundamentals of Biochemistry Introduction The elucidation of the structure of the double helix by James Watson and Francis Crick in 1953 provided a hint as to how DNA is copied during the process of DNA replication. Separating the strands of the double helix would provide two templates for the synthesis of new complementary strands, but exactly how new DNA molecules were constructed was still unclear. In one model, semiconservative replication, the two strands of the double helix separate during DNA replication, and each strand serves as a template from which the new complementary strand is copied. After replication in this model, each double-stranded DNA includes one parental or “old” strand and one daughter or “new” strand. There were two competing models also suggested: conservative and dispersive, which are shown in Figure \(1\). Matthew Meselson and Franklin Stahl devised an experiment in 1958 to test which of these models correctly represents DNA replication, as shown in Figure \(2\). They grew the bacterium, Escherichia coli for several generations in a medium containing a “heavy” isotope of nitrogen (15N) that was incorporated into nitrogenous bases and, eventually, into the DNA. This labeled the parental DNA. The E. coli culture was then shifted into a medium containing 14N and allowed to grow for one generation. The cells were harvested and the DNA was isolated. The DNA was separated by ultracentrifugation, during which the DNA formed bands according to its density. DNA grown in 15N would be expected to form a band at a higher density position than that grown in 14N. Meselson and Stahl noted that after one generation of growth in 14N, the single band observed was intermediate in position in between DNA of cells grown exclusively in 15N or 14N. This suggested either a semiconservative or dispersive mode of replication. Some cells were allowed to grow for one more generation in 14N and spun again. The DNA harvested from cells grown for two generations in 14N formed two bands: one DNA band was at the intermediate position between 15N and 14N, and the other corresponded to the band of 14N DNA. These results could only be explained if DNA replicates in a semiconservative manner. Therefore, the other two models were ruled out. As a result of this experiment, we now know that during DNA replication, each of the two strands that make up the double helix serves as a template from which new strands are copied. The new strand will be complementary to the parental or “old” strand. The resulting DNA molecules have the same sequence and are divided equally into the two daughter cells. Think about It: What would have been the conclusion of the Meselson-Stahl experiment if, after the first generation, they had found two bands of DNA? To synthesize double-stranded DNA, the parental strands must separate so DNA polymerases can copy both strands. As all DNA polymerases synthesize new DNA in a 5' to 3' direction from a 3' to 5' template, different mechanisms are used to faithfully synthesize both parental strands. The general mechanism is shown in Figure \(3\). Figure \(3\): The replication fork. Leading-strand synthesis proceeds continuously in the 5' to 3' direction. Lagging-strand synthesis also occurs in the 5' to 3' direction, but in a discontinuous manner. An RNA/DNA primer (labeled in green) initiates leading-strand synthesis and every Okazaki fragment on the lagging strand. Small RNA primers are needed for the new strands. Short (1000-2000 NT) DNA (Okazaki) fragments are made on the 3'-5' parental strand. Ultimately the RNA primers are degraded and filled, and the Okazaki fragments ligated. We will discuss replication in detail for E. Coli, a model prokaryote, followed by replication in eukaryotes. Figure \(4\) shows a general overview of a DNA "replication fork" from where DNA strand synthesis proceeds.. DNA Replication in E. Coli DNA replication has been well studied in bacteria primarily because of the small size of the genome and the mutants that are available. E. coli has 4.6 million base pairs (Mbp) in a single circular chromosome and all of it is replicated in approximately 42 minutes, starting from a single origin of replication and proceeding around the circle bidirectionally (i.e., in both directions), as shown in Figure \(5\). This means that approximately 1000 nucleotides are added per second. The process is quite rapid and occurs with few errors. E. coli has a single origin of replication, called oriC, on its one chromosome. The origin of replication is approximately 245 base pairs long and is rich in adenine-thymine (AT) sequences. Replication Overview - E. Coli The open regions of DNA that are actively undergoing replication are called replication forks. All the proteins involved in DNA replication aggregate at the replication forks to form a replication complex called a replisome. The initial assembly of the complex that initiates primer synthesis is called the primosome. Table \(1\) below show the components that assemble at the replication fork to form the E. Coli replisome. Table \(1\): Enzymes involved in DNA Replication in the prokaryote, E. coli In E. coli, DNA replication is initiated at the single origin of replication, oriC. Binding of the initiator protein, DnaA, locally unfolds the DNA to form two template ssDNA, which bind DnaB helicase. A DnaB hexamer adds to each strand in a process promoted by DnaC, a helicase loader. The single-stranded DNA binding protein B (SSPB)binds to and protects the rest of the ssDNA, preventing further binding by DnaB. The primase, DnaG, is recruited to the site by the DnaB hexamer and synthesizes the RNA primers. DnaB also recruits DNA polymer III holoenzyme (PolIII HE) which binds through a β clamp. All of the bound proteins collectively form the replisome. An overview of E. Coli replisome is shown in Figure \(6\). Figure \(6\): The bacterial replisome. Ilic, S.; Cohen, S.; Singh, M.; Tam, B.; Dayan, A.; Akabayov, B. DnaG Primase—A Target for the Development of Novel Antibacterial Agents. Antibiotics 2018, 7, 72. https://doi.org/10.3390/antibiotics7030072 Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/) Once assembled, replisomes move in opposite directions from the single oriC in the E. Coli chromosome. They meet at the opposite ends at a termination site (ter) to which Tus proteins are bound that create ‘replication fork traps'. After completion of DNA replication, the newly synthesized genomes are separated and segregated to daughter cells. An alternative term, the primosomes, is used to describe a subcomplex of the replisome which starts replication of the E. Coli chromosome, as well as some phages and plasmids. It contains 6 proteins including helicases and primases, and catalyzes the movement of the replication form by unwinding and primer synthesis. The motor protein helicases, use ATP to move along the ds-DNA backbone, unraveling it as it proceeds. The human genome has genes for 64 RNA and 31 DNA helicases (about 1% of eukaryotic genes). Primase and Polymerase activities The synthesis of both RNA strands by the DnaG primase, and DNA strands by DNA polymerase III holoenzyme (pol III) occurs at each start site for an Okazaki fragment. Both enzymes bind to the conserved carboxy-terminal tail of the single-stranded DNA-binding protein (SSB). It turns out that they can both be bound simultaneously. The primase (DnaG) has three domains: • N-terminus that binds the template • RNA polymerase domain • C-terminus that binds helicase and the C-terminus of SSB. Primase is displaced by polIII after about 10 nucleotides have been added to the RNA primer so DNA synthesis can now occur at the 3' end of the primer. Figure \(7\) shows how the primase to polymerase switch is made. Figure \(7\): Schematic representation of the primase-to-polymerase switch during DNA replication in E. coli. Bogutzki, A., Naue, N., Litz, L. et al. Sci Rep 9, 14460 (2019). https://doi.org/10.1038/s41598-019-51031-0. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ Panel (a): Two primase molecules cooperate in the synthesis of the RNA primer (red). Panel (b): For elongation of the primer, pol III enters the complex, whereupon primase and pol III are concurrently bound to the primed site, possibly via interactions with the C-termini of an adjacent SSB tetramer. Panel (c): During pol III-mediated elongation of the primer by several nucleotides, both enzymes stay bound to the template. Only after the primer has been elongated by more than 10 nucleotides, one of the primases is released in the G4ori system. It is presumably the displacement of SSB by pol III that causes the consequent dissociation of primase. Whereas in the G4ori system, the two primases are positioned by hairpin structures that prevent SSB from binding to this part of the origin.  At the E. coli replication fork, primases are brought into contact via their interaction with the replicative helicase DnaB. What happens if the replication fork does not move to the termination site? If the DNA is damaged or if the replisome falls off of the chromosome, it can rebind and restart using Pri proteins. PriA is a DNA helicase that can bind to replication forks through DNA motifs and through interactions with SSBs. Other proteins involved include PriB, PriC, DnaT, DnaC, DnaB helicase, and DnaG primase as illustrated in Figure \(8\). The primosome and replisome are complicated in structure and in their functional activity. Words go only so far in painting an image of how it works. To help we show a few different images of the replisome of E. Coli below. The first is shown in Figure \(9\). In this diagram, the leading strand is shown in the upper right end of the diagram. The central bottom loop shows the lagging strand. The τ3δδ′ψχ is the clamp loader and the DnaB (red) is hexameric. The ssDNA in the lagging strand loop is bound by ssDNA binding proteins (SSB). Figure \(10\) shows the rebind primosome which is mostly similar to the regular one. Figure \(10\): Mechanisms of helicase loading leading to replisome assembly in E. coli. (A)Recognition and melting of the oriC locus during initiation by DnaA. (B)Recognition of abandoned fork structures during replisome reloading by PriA and PriC. All pathways converge on the loading of the replicative helicase DnaB, which acts as an assembly platform for the remaining replisome components. Finally, Figure \(11\) shows models of DNA polymerase for lagging strand synthesis. Figure \(11\): Usage of DNA polymerase during lagging strand synthesis. (A)S chematic of the E. coli replisome during the elongation step of an Okazaki fragment. (B )Lagging strand polymerase meets the RNA primer of the previous Okazaki fragment and stops synthesis. (C) Current model of events following completion of an Okazaki fragment. DNA polymerase is released from the β clamp (step 1) and the same molecule rebinds to a new β clamp to start the next Okazaki fragment (step 2). (D)An alternative model based on evidence from T4 and T7 replisomes. After completing the Okazaki fragment, the DNA polymerase detaches from the rest of the replisome (step 1). A new molecule of DNA polymerase is recruited to the replisome (step 2) and engages in the synthesis of a new Okazaki fragment. In this tentative model, a local pool of “spare” polymerases may facilitate their exchange and additional components may exchange along with the polymerase (not depicted) E. Coli DNA Polymerases E. Coli has 5 DNA polymerases. DNA polymerase I aids in lagging strand synthesis as it removes the RNA primers and incorporates DNA in its place. DNA polymerase II, may play an editing role following lagging strand synthesis by DNA polymerase I. DNA polymerases I and II also play a role in DNA repair, as do DNA polymerases IV and V. DNA polymerases are shaped like a right hand in overall shape with three domains named palm, fingers, and thumb. The bottom of the cleft formed by the three domains forms the polymerase active site in the Palm domain, The monomeric nucleotides to be added bind through the finger domain, while the thumb domains facilitates the dissociation of the newly synthesized DNA. These features are illustrated for a polymerase that requires host thioredoxin for a bacteriophage T7 DNA polymerase in Figure \(12\). Figure \(12\): Structure of T7 DNA replication complex. Melum 103 - Own work, CC BY-SA 4.0, https://commons.wikimedia.org/w/inde...curid=38408627 DNA Polymerase III Pol III is a fascinating enzyme. It consists of an αεθ core with both 5'-3' polymerase and 3′−5′ proofreading activities, a β2 ring-shaped "sliding clamp" that keeps the enzyme on the DNA track (processive) without iteratively jumping off and rebinding (distributive), and a (τ/γ)3δδ′ψχ clamp loader. The SSB protein has a conserved amphiphilic C-terminus that binds both DnaG (primase) and the χ subunit of the clamp loader. A After primer addition by DnaG, the β2clamp of polIII is brought to the end of the primer terminus by the clamp loader, after which α and ε subunits bind the clamp. The holoenzyme can add ∼1000 Nt/s and over 150 kb without falling off. Hence it is a very processive enzyme. Figure \(13\) shows an interactive iCn3D model of the the E. coli replicative DNA polymerase III (alpha, beta2, epsilon, tau complex) bound to DNA (5FKV) • N-terminal domains of α (αNTD, residues 1–963, are colored in salmon • OB (964–1072) on the C-term domain of α (αCTD) colored brown, • τ-binding domains (TBD, 1173–1160) on the C-term domain of α (αCTD) colored dark salmon, • ε in yellow • θ in orange (?) • β2 in aquamarine • τC in slate gray • DNA in spacefill, backbone magenta and purple, bases in CPK colors Overall, there are significant conformational changes in the DNA Polymerase III complex upon binding to the DNA that cause the tail of the polymerase to move from interacting with the clamp in the DNA-bound state to a position 35 Å away from the clamp in the DNA-free state. It has been hypothesized that this large conformational change may help the polymerase act as a switch to facilitate the lagging strand synthesis. On the lagging strand, the polymerase repositions to a newly primed site every ∼1000 bp. To do so, the polymerase needs to release both clamp and DNA. The switch-like movement of the polymerase tail may play a part in the release and consequent repositioning of the polymerase at the end of the Okazaki fragment. Video 25.1.1: DNA Binding Induces Large Conformational Changes in the DNA Polymerase III Complex(click link to view). The video shows the linear morphing of the DNA-free to the DNA-bound state showing the large conformation change between the two states. The green subunit is the β-clamp, The α-subunit is shown in orange with the active-site residues in magenta, the α-C-terminal domain (α-CTD shown in brown, the ε-subunit in yellow, and the τ-tail shown in blue. Video from: Fernandez-Liero, R., et al. (2015) eLife 4:e11134 The complex can also proofread the newly synthesized DNA. This requires some conformational changes in the polIII complex, including a rotation/tilt of the dsDNA against the β2 ring-shaped "sliding clamp". The thumb domain moves between the two DNA strands containing a mismatch and produced a distorted DNA. The epsilon subunit, a nuclease, can reach the mismatched nucleotide and clip it off. Figure \(14\) shows an interactive iCn3D model of the E. coli replicative DNA polymerase III-clamp-exonuclease-theta complex bound to DNA in the editing mode (5M1S) • PolIII α brown, • PolIII ε in yellow • PolIII θ in orange • PolIII β2 in cyan • DNA in spacefill, backbone primer in magenta, the template in purple, and bases in CPK colors DNA Polymerase I DNA polymerase I, as does polIII, has a 5' to 3' polymerase activity. Also, both have a 3' to 5' exonuclease activity for proofreading as well as a 5'-3' exonuclease to remove RNA primers. It contains three domains, a 5'-3' exonuclease followed by a 3'-5' exonuclease, then the polymerase domain. Selective proteolysis between the first two domains produces the Klenow fragment. In contrast, the 5'-3' exonuclease of polIII is in the separate epsilon subunit. Figure \(15\) shows an interactive iCn3D model of the predicted AlphaFold structure of E. Coli DNA Polymerase I (P00582) https://structure.ncbi.nlm.nih.gov/i...oNSC2GJAsMEUk6 • 5' to 3' exonuclease, 1-323, magenta • 3' to 5' exonuclease, 324-517, orange • 5' to 3' polymerase, 521-928, cyan • Val700-Arg713, Motif A, yellow • Klenow fragment: 324-928 Key aspartate and glutamates involved in the polymerase active site are shown in sicks and labeled. Motif A is conserved in prokaryotic DNA polymerases. Essential roles of motif A in catalysis include interaction with the incoming dNTP and coordination with two divalent metal ions that are required for the polymerization reaction. Note the distance between the 3' to 5' exonuclease and the 5'-3' polymerase. Other enzyme activities DNA Ligase DNA Ligase enzymes seal the breaks in the backbone of DNA that are caused during DNA replication, DNA damage, or during the DNA repair process. The biochemical activity of DNA ligases results in the sealing of breaks between 5′-phosphate and 3′-hydroxyl termini within a strand of DNA. DNA ligases have been differentiated as being ATP-dependent or NAD+-dependent depending on the co-factor (or co-substrate) that is used during their reaction. Typically, more than one type of DNA ligase is found within an organism. Figure \(16\) shows the structure of E. coli LigA in complex with nicked adenylated DNA from PDB 2OWO, visualized by UCSF Chimera. The various domains are indicated by different colors and relate to Pfam domains indicated. Figure \(16\): Structure of DNA ligase. Pergolizzi, G., Wagner, G.K, and Bowater, R.P. (2016) Biosci Rep 36(5) e00391 DNA ligase enzyme is covalently modified by the addition of the AMP moiety to a Lysine residue on the enzyme. The AMP derives from the ATP or NADH cofactor. The downstream 5'-phosphate at the site of the DNA nick is able to mediate a nucleophilic attack on the AMP-enzyme complex, causing the AMP to transfer to the 5'-phosphate position of the DNA. The AMP serves as a good leaving group for the nucleophilic attack of the upstream 3'-OH with the 5'-phosphate to seal the DNA backbone, and release the AMP. DNA ligase can use either adenosine triphosphate (ATP) or nicotinamide adenine dinucleotide (NAD+) as a cofactor. Figure \(17\) shows a mechanism of the ligation reaction, which is powered by ATP hydrolysis. Figure \(18\) shows an interactive iCn3D model of Human DNA Ligase I bound to 5'-adenylated, nicked DNA (1X9N) The protein has three domains: • DNA binding domain (DBD), shown in magenta. In contrast to most DNA binding proteins, Ligase I binds to the minor grove around the area of DNA damage. • Adenylation domain (AdD), shown in cyan, has covalently attached cofactor AMP and key catalytic residues shown as sticks and labeled. It ligates the broken DNA and forms a phosphodiester bond. • The OB-fold domain (OBD), shown in yellow, facilitates catalysis as it binds and unwinds over the short region. The DNA strands are as follows: • ss DNA terminated with dideoxy is shown in green • ss template DNA is shown in brown • ss 5'-phosphorylated DNA is shown in gray The AdD and OBD domains are similar in structure to other covalent nucleotidyltransferases involved in DNA and RNA ligation and capping of messenger RNA. Glu 566, Glu 621, and Arg 573 interact with AMP and probably help determine the specificity of the AMP cofactor (over GTP). Divalent cations are required for catalysis but are not present in the above structure. A E566K mutation leads to severe immunodeficiency. Lys 568 forms the covalent AMP adduct. The dideoxynucleoside in the structure is not optimally positioned for reaction with 5'P of AppDNA. Glu 720 and Glu 621 are highly conserved and presumably involved in metal ion binding. Zoom into the structure to observe the changes at the 3' end of the dideoxy DNA and 5' end of the phosphorylated DNA. These nucleotides are given the abbreviation X in iCn3D as they are modified. Topoisomerases We just studied these in a previous section but here is a review for this section. The unwinding of the double-stranded helix at the replication fork generates winding tension in the DNA in the form of positive supercoils upstream of the replication fork. Enzymes called topoisomerases counteract this by introducing negative supercoils into the DNA in order to relieve this stress in the helical molecule during replication. There are four known topoisomerase enzymes found in E. coli that fall into two major classes, Type I Topoisomerases and Type II Topoisomerases, as shown in Figure \(19\). Topoisomerase I and III are Type I topoisomerases, whereas DNA gyrase and Topoisomerase IV are Type II topoisomerases. Goodsell, D.S. (2015) RCSD PDB-101 Molecule of the Month Type I Topoisimerase Type I Topoisomerases relieve tension caused during the winding and unwinding of DNA. One way that they can do this is by making a cut or nick in one strand of the DNA double helix as shown in Figure \(19\). The 5'-phosphoryl side of the nicked DNA strand remains covalently bound to the enzyme at a tyrosine residue, while the 3'-end is held noncovalently by the enzyme. The Type I topoisomerases rotate or spin the 3'-end of the DNA around the intact DNA strand. This releases the overwinding in the DNA and effectively releases tension. The enzyme completes the reaction by resealing the phosphodiester backbone or ligating the broken strand back together. Overall, only one strand of the DNA is broken during the reaction mechanism and there is NO requirement of ATP during the reaction. The E. coli Topo I enzyme can only remove negative DNA supercoils, but not positive ones. Thus, this enzyme is not involved in relieving the positive supercoiling caused by the DNA helicase during replication. This is in contrast to eukaryotic Topo I that can relieve both positive and negative supercoiling. Although E. coli Topoisomerase I is not directly involved in relieving the tension caused by DNA replication, it is essential for E. coli viability. It is thought to help balance the actions of the Type II topoisomerases and help maintain optimal supercoiling density within the chromosomal DNA. Thus, Topo I is thought to help maintain the homeostatic balance of chromosome supercoiling within E. coli. Topo III, which is also a Type I Topoisomerase, appears to play a role in the decatenation of the daughter chromosomes during DNA replication, but does not play a role in the relaxation of supercoiling. Type II Topoisomerases have multiple functions within the cell. They can increase or decrease winding tension within the DNA or they can unknot or decatanate DNA that has become tangled with another strand as shown in Figure \(20\). It does so by a more dangerous method than their Type I counterparts, by breaking both strands of the DNA during their reaction mechanism. The enzyme is covalently attached to both broken sides while the other DNA helix is passed through the break. The double-stranded break is then resealed. The proposed type II topoisomerase reaction cycle is exemplified by topoisomerase IV. Topoisomerase IV subunits are denoted in grey, cyan, and yellow. The gate or G-DNA is in green and the transported or T-DNA is in mauve. ATP bound to the ATPase domains is denoted by a red dot. In step 1, the G-DNA binds with the enzyme. ATP and the T-DNA segment associated with the enzyme in step 2. In step 3, the G-DNA is cleaved and the T-DNA is passed through the break. Drug-targetable domains within the type II topoisomerase complex are highlighted in subsections A, B, and C with examples on the right-hand side of the figure. Type II Topoisomerase - D DNA gyrase is the type II topoisomerase enzyme that is primarily involved in relieving positive supercoiling tension that results due to the helicase unwinding at the replication fork. Type II Topoisomerases, especially Topo IV, also address a key mechanistic challenge that faces the bacterial replisome during the termination of DNA replication. The circular nature of the bacterial chromosome dictates that a pair of replisomes that initiate from a single origin of replication will eventually converge on each other in a head-to-head orientation. Positive supercoiling accumulates between the the two replisomes as they converge, but the activity of DNA gyrase, which normally removes positive supercoils, becomes limited by the decreasing amount of template DNA available. Instead, supercoils may diffuse behind the replisomes, forming precatenanes between newly replicated DNA; in E.coli these must be resolved by Topo IV for chromosome segregation to occur. Termination of Replication If starting replication is critically important and obviously highly controlled as illustrated above, then termination of replication must be equally critical, otherwise genome instability would arise. There is one discrete origin of replication in E. Coli, oriC, with a defined sequence. In contrast, there are 10, 23 base-pair, nonpalindromic termination sites (Ter)of slightly different sequences. These bind the termination protein, Tus. The affinity of Tus for the Ter site depends on the Ter sequence and, in general,is tight with a KD in the picomolar range. There are two types of Ter-Tus complexes, one an open"permissive" conformation that allows replication to continue, and a locked, "nonpermissive" form that stops it. In the nonpermissive conformation, a key and conserved cytosine on the leading strand at a conserved GC base pair is flipped out into a cytosine binding pocket, which you can think of as a "stop sign" for replication. If you think of the E. Coli circular chromosome as a clock with the oriC at 12 o'clock, there are 5 Ter sequences as the replication fork move counter-clockwise at about 7 o'clock and another 5 as the fork move clockwise at around 5 o'clock. The sequences run in opposite polarity to prevent the left-side replication fork from entering the right-hand side as it moves around the chromosome and vice versa. The replisome displaces the Tus proteins at the permissive Ter sites but stops at the nonpermissive site, where DnaB helicase unwinds the DNA, flipping out the cytosine as the locked conformation forms. These processes are illustrated in Figure \(21\). Panel A, E. coli contains a single circular chromosome, which replicates bidirectionally from a single origin (small oval). The direction of replisome travel from the origin is depicted by arrows. Chromosomal midpoint indicated by a straight line. The location of the ter sites on the E. coli chromosome is shown relative to oriC. Permissive orientation is displayed in light blue, nonpermissive orientation is displayed in dark blue. Panel B, the structure of Tus-ter (PDB ID: 2I06) illustrating the nonpermissive and permissive faces (left) and the “locked” conformation formed by DNA unwinding at the nonpermissive face (right). The cytosine base at position 6 of ter (C6), which flips into a specific binding site on the nonpermissive face of Tus to form the “lock,” is indicated. Replication can proceed at the Ter site if the replisome is moving from the light blue to dark blue sequences of the TER site. The TER site hence exhibits polarity. Panel A, E. coli contains a single circular chromosome, which replicates bidirectionally from a single origin (small oval). The direction of replisome travel from the origin is depicted by arrows. Chromosomal midpoint indicated by a straight line. The location of ter sites on the E. coli chromosome is shown relative to oriC. Permissive orientation is displayed in light blue, nonpermissive orientation is displayed in dark blue. Panel B, structure of Tus-ter (PDB ID: 2I06) illustrating the nonpermissive and permissive faces (left) and the “locked” conformation formed by DNA unwinding at the nonpermissive face (right). The cytosine base at position 6 of ter (C6), which flips into a specific binding site on the nonpermissive face of Tus to form the “lock,” is indicated. Figure \(22\) Crystal structure (PDB code: 2I06) of the locked Tus–Ter complex shows the flipped C(6) base at the non-permissive face (5) Figure \(22\): Crystal structure (PDB code: 2I06) of the locked Tus–Ter complex shows the flipped C(6) base at the non-permissive face (5). Pandey et al. 2015 Jul 13; 43(12): 5924–5935. doi: 10.1093/nar/gkv527. Creative Commons Attribution License (http://creativecommons.org/licenses/by-nc/4.0/), Figure \(23\) shows interactive iCn3D models of the Escherichia Coli Replication Terminator Protein (Tus) Complexed With TerA DNA in open (left) or locked form (right). Escherichia Coli Replication Terminator Protein (Tus) Complexed With TerA DNA (2I05) Escherichia Coli Replication Terminator Protein (Tus) Complexed With DNA- Locked form (2I06) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?7mc7UTqLBbgRZP8p8 (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?bScTvhNGAMD456vt9 In the figures, the C6 that is flipped out in the locked nonpermissive form (right) is shown in spacefill and labeled as C324. The Arg 198 alters its orientation to allow the C6 to flip out and form the locked conformational form. A summary of the process of DNA replication is shown in Video 25.1.2 Click Here to View Video Video 9.2 Overview of the DNA Replication Process DNA Replication of Extrachromosomal Elements: Plasmids and Viruses To copy their nucleic acids, plasmids and viruses frequently use variations on the pattern of DNA replication described for prokaryote genomes. We will focus here on one style known as the rolling circle method. Whereas many bacterial plasmids replicate by a process similar to that used to copy the bacterial chromosome, other plasmids, several bacteriophages, and some viruses of eukaryotes use rolling circle replication as shown in Figure \(24\). Figure \(24\): Rolling Circle Replication. The process of rolling circle replication is initiated by a single stranded nick in the DNA. Within prokaryotes, DNA polymerase III is utilized to generate the daughter strand. DNA ligase rejoins nicks in the backbone and enables the initiation of DNA synthesis of the second daughter strand. Figure by Parker, N., et.al. (2019) Openstax. The circular nature of plasmids and the circularization of some viral genomes on infection make this possible. Rolling circle replication begins with the enzymatic nicking of one strand of the double-stranded circular molecule at the double-stranded origin (dso) site. In bacteria, DNA polymerase III binds to the 3′-OH group of the nicked strand and begins to unidirectionally replicate the DNA using the un-nicked strand as a template, displacing the nicked strand as it does so. Completion of DNA replication at the site of the original nick results in the full displacement of the nicked strand, which may then recircularize into a single-stranded DNA molecule. RNA primase then synthesizes a primer to initiate DNA replication at the single-stranded origin (sso) site of the single-stranded DNA (ssDNA) molecule, resulting in a double-stranded DNA (dsDNA) molecule identical to the other circular DNA molecule. DNA Replication in Eukaryotes The Cell Cycle The cell cycle is an ordered series of events involving cell growth and cell division that produces two new daughter cells. Cells on the path to cell division proceed through a series of precisely timed and carefully regulated stages of growth, DNA replication, and division that produce two genetically identical cells. The cell cycle has two major phases, interphaseand the mitotic phase, as shown in Figure \(25\). During interphase, the cell grows and DNA is replicated. During the mitotic phase, the replicated DNA and cytoplasmic contents are separated and the cell divides. Watch this video about the cell cycle: http://openstax.org/l/biocellcyc Figure \(25\): Diagram of the Cell Cycle. Fowler, S., et.al. (2013) Openstax A cell moves through a series of phases in an orderly manner. During interphase, G1 involves cell growth and protein synthesis, the S phase involves DNA replication and the replication of the centrosome, and G2 involves further growth and protein synthesis. The mitotic phase follows interphase. Mitosis is nuclear division during which duplicated chromosomes are segregated and distributed into daughter nuclei. Usually, the cell will divide after mitosis in a process called cytokinesis in which the cytoplasm is divided and two daughter cells are formed. During interphase, the cell undergoes normal processes while also preparing for cell division. For a cell to move from interphase to the mitotic phase, many internal and external conditions must be met. The three stages of interphase are called G1, S, and G2. The first stage of interphase is called the G1 phase, or first gap, because little change is visible. However, during the G1 stage, the cell is quite active at the biochemical level. The cell is accumulating the building blocks of chromosomal DNA and the associated proteins, as well as accumulating enough energy reserves to complete the task of replicating each chromosome in the nucleus. Throughout interphase, nuclear DNA remains in a semi-condensed chromatin configuration. In the S phase (synthesis phase), DNA replication results in the formation of two identical copies of each chromosome—sister chromatids—that are firmly attached at the centromere region,as shown in Figure \(26\). At this stage, each chromosome is made of two sister chromatids and is a duplicated chromosome. The centrosome is duplicated during the S phase. The two centrosomes will give rise to the mitotic spindle, the apparatus that orchestrates the movement of chromosomes during mitosis. In mammals, the centrosome consists of a pair of rod-like centrioles at right angles to each other. Centrioles help organize cell division. Centrioles are not present in the centrosomes of many eukaryotic species, such as plants and most fungi. Figure \(26\): Human Chromosome Structure. Figure A from: The National Human Genome Research Institute, and Figure B from: The School of Biomedical Sciences Wiki (A) Shows a spectral karyogram of a normal human female. Humans have a total of 23 pairs of chromosomes for a total of 46. Each pair of chromosomes are referred to as homologous chromosomes as they contain copies of the same gene regions. Each of the homologous pairs of chromosomes is stained the same color. Chromosomes are shown in their condensed, unreplicated state. (B) Shows a schematic diagram of a single chromosome before (lower diagram) and after (upper diagram) replication. Upon replication, the identical copies of the chromosome are called sister chromatids and are linked together at the centromere structure. Figure A from: The National Human Genome Research Institute, and Figure B from: The School of Biomedical Sciences Wiki In the G2 phase, or second gap, the cell replenishes its energy stores and synthesizes the proteins necessary for chromosome manipulation. Some cell organelles are duplicated, and the cytoskeleton is dismantled to provide resources for the mitotic spindle. There may be additional cell growth during G2. The final preparations for the mitotic phase must be completed before the cell is able to enter the first stage of mitosis. To make two daughter cells, the contents of the nucleus and the cytoplasm must be divided. The mitotic phase is a multistep process during which the duplicated chromosomes are aligned, separated, and moved to opposite poles of the cell, and then the cell is divided into two new identical daughter cells. The first portion of the mitotic phase, mitosis, is composed of five stages, which accomplish nuclear division. The second portion of the mitotic phase, called cytokinesis, is the physical separation of the cytoplasmic components into two daughter cells. If cells are not traversing through one of the phases of interphase or mitosis, they are said to be in G0 or a resting state. If cells enter G0 permanently, they are said to have entered a stage of replicative senescence and will no longer be maintained for long-term viability within the organism. The progression of cells through the cell cycle requires the coordinated actions of specific protein kinases, known as cyclin-dependent kinases. Cyclin-dependent kinases are usually abbreviated as CDK or CDC proteins. CDK/CDC proteins require the binding of a regulatory cyclin protein to become activated, as shown in Figure \(27\). The major cyclin proteins that drive the cell cycle in the forward direction, are expressed only at discrete times during the cell cycle. When activated by a cyclin counterpart, CDK/CDC enzymes phosphorylate downstream targets involved with cell cycle progression. For example, the primary cyclin-CDK complex involved in the initiation of DNA replication during S-phase is the CyclinE-CDK2 complex. CDK2 is activated by the expression and binding of Cyclin E during late G1 phase. This causes CDK2 to phosphorylate downstream targets, including the retinoblastoma tumor suppressor protein, pRb. pRB normally binds and inhibits the activity of transcription factors from the E2F family. Following the release of E2F transcription factors from pRb, E2Fs activate the transcription of genes involved in DNA replication and the progression of cells into S-phase. Panel (A) shows CDK-cyclin complexes with direct functions in regulating the cell cycle are shown. CDK3/cyclin C drives cell cycle entry from G0. CDK4/6/cyclin D complexes initiate phosphorylation of the retinoblastoma protein (pRb) and promote the activation of CDK2/cyclin E complex. In late G1, CDK2/cyclin E complex completes phosphorylation and inactivation of pRb, which releases the E2F transcription factors and G1/S transition takes place. DNA replication takes place in S phase. CDK2/cyclin A complex regulates progression through S phase and CDK1/cyclin A complex through G2 phase in preparation for mitosis (M). Mitosis is initiated by CDK1/cyclin B complex. Panel (B) Shows the cyclical nature of cyclin expression during cell cycle progression. Cyclin abundance is regulated by transcriptional expression and rapid protein degradation. Thus, their biological activity is targeted at very specific time points during the cell cycle progression. Replication Initiation Origin organization, specification, and activation in eukaryotes are more complex than in bacterial or archaeal kingdoms and significantly deviate from the paradigm established for prokaryotic replication initiation. The large genome sizes of eukaryotic cells, which range from 12 Mbp in S. cerevisiae to 3 Gbp in humans, necessitates that DNA replication starts at several hundred (in budding yeast) to tens of thousands (in humans) origins to complete DNA replication of all chromosomes during each cell cycle, as shown in Figure \(27\). Figure \(27\): Eukaryotic chromosomes are typically linear, and each contains multiple origins of replication. The top figure is a graphic representation of the eukaryotic origins of replication, while the bottom image is a Cryo-electron micrograph image. The figure on the top is from Parker, N. et al. and the figure on the bottom is from Fritensky, B. and Brien, N With the exception of S.cerevisiae and related Saccharomycotina species, eukaryotic origins do not contain consensus DNA sequence elements but their location is influenced by contextual cues such as local DNA topology, DNA structural features, and chromatin environment. Nonetheless, eukaryotic origin function still relies on a conserved initiator protein complex to load replicative helicases onto DNA during the late M and G1 phases of the cell cycle, a step known as origin licensing. In contrast to their bacterial counterparts, replicative helicases in eukaryotes are loaded onto origin duplex DNA in an inactive, double-hexameric form and only a subset of them (10–20% in mammalian cells) is activated during any given S phase, events that are referred to as origin firing. The location of active eukaryotic origins is therefore determined on at least two different levels, origin licensing to mark all potential origins, and origin firing to select a subset that permits assembly of the replication machinery and initiation of DNA synthesis. The extra licensed origins serve as backup and are activated only upon slowing or stalling of nearby replication forks, ensuring that DNA replication can be completed when cells encounter replication stress. Together, the excess of licensed origins and the tight cell cycle control of origin licensing and firing embody two important strategies to prevent under- and overreplication and to maintain the integrity of eukaryotic genomes. Human Primosome In humans,, the primosome contains primase and DNA polymerase α (Polα), and makes RNA-DNA primers to which deoxynucleotides are added by polymerases δ and ϵ. Hence there are two catalytic sites for addition or ribo- and deoxyribonucleotides. The structure of the human primosome and the C-terminal domain of the primase large subunit (p58C) with bound DNA/RNA duplex is presented below. p58C coordinates the catalytic activities. As with other polymerases, primase synthesis of RNA primers has the following steps: • initiation (rate limiting): primase binds to DNA and makes a dinucleotide RNA; • elongation, which is not as fast as DNA replication since it is less processive, adding only around 10 nucleotides. These short fragments are moved to Polα where deoxynucleotides are added with inactivation of the primase • termination. The structures of the enzymes are as follows: Human Polα consists of a : • large catalytic subunit (p180) with a C-terminal p180C domain with two Zn2+ binding modules. • smaller accessory subunit (p70) with an N-terminal (p70N), a phosphodiesterase, and oligonucleotide/oligosaccharide-binding (OB) domains. Human primase consist of • catalytic (p49) • regulatory (p58) subunits with two domains, the N-terminal (p58N), which interacts with p49 and which connects primase and Polα, and a C-terminal (p58C) which contains an iron-sulfur cluster involved in substrate binding and primase activity. The structures are shown in Figure \(28\). Figure \(28\): Structure of the human primosome hetero-tetramer complex. Baranovskiy et al. JBC, 291, 10006-10020 (2016). DOI: https://doi.org/10.1074/jbc.M116.717405. Creative Commons Attribution (CC BY 4.0) Panel A shows a schematic representation of the domain organization. The flexibly tethered domains are shown as separate parts. p58C coordinates the iron-sulfur cluster. Exo* is an exonuclease domain with no associated activity due to the evolutionary substitution of the catalytic amino acid residues; PDE, phosphodiesterase. Pane B shows the crystal structure of the primosome. Subunits are shown as schematics and colored as in A. The α-carbons of catalytic aspartates are shown as purple spheres. Figure \(29\) shows an interactive iCn3D model of the Human primosome without nucleic acids (5EXR) DNA primase small: catalytic (p49) - dark green DNA primase large: regulatory (p58) subunits. p58 has two distinct domains, N-terminal (p58N light blue) and C-terminal (p58C gray/purple), connected with an 18-residue linker (253–270) . p58N interacts with p49 and connects primase with Polα ), and an iron-sulfur cluster containing p58C plays an important role in substrate binding and primase activity DNA polymerase alpha catalytic subunit: large catalytic subunit (p180). has p180core (orange) and linker 1251-1265 then the C-terminal domain (p180C - blue) connects to small subunit p70. (p180C) contains Zn1 and Zn2 bind site DNA polymerase alpha subunit B: smaller accessory subunit (p70) with 3 domains: p70N (light green) then linker 79-156 BOTH NOT SHOWN IN STRUCTURE), the P70 phosphodiesterase, and oligonucleotide/oligosaccharide-binding (OB) domains (combined magenta). Figure \(30\) shows an interactive iCn3D model of the C-terminal domain of the human DNA primase large subunit with bound DNA template/RNA primer (5F0Q) The ss-DNA is shown with a pale green backbone while the RNA backbone is shown in magenta. The FeS cluster and a Mg2+ ion are shown in the catalytic subunit. The Mg2+ is shown interacting with a terminal GTP of the RNA. Figure \(31\) shows an interactive iCn3D model of the catalytic core of human DNA polymerase alpha in a ternary complex with an RNA-primed DNA template and dCTP (4QCL) The ss-DNA is shown with a cyan backbone while the RNA primer backbone is shown in magenta. dCTP is shown in spacefill. Eukaryotic DNA polymerases Similar to DNA replication in prokaryotes, DNA replication in eukaryotes occurs in opposite directions between the two new strands at the replication fork. Within eukaryotes, two replicative polymerases synthesize DNA in opposing orientations, as shown in Figure \(32\). Polymerase ε (epsilon) synthesizes DNA in a continuous fashion, as it is “pointed” in the same direction as DNA unwinding. Similar to bacterial replication, this strand is known as the leading strand. In contrast, polymerase δ (delta) synthesizes DNA on the opposite template strand in a fragmented, or discontinuous, manner and this strand is termed the lagging strand. The discontinuous stretches of DNA replication products on the lagging strand are known as Okazaki fragments and are about 100 to 200 bases in length at eukaryotic replication forks. Owing to the “lagging” nature, the lagging strand generally contains a longer stretch of ssDNA that is coated by single-stranded binding proteins, which stabilizes ssDNA templates by preventing secondary structure formation or other transactions at the exposed ssDNA. In eukaryotes, ssDNA stabilization is maintained by the heterotrimeric complex known as replication protein A (RPA) (Figure 9.19). Each Okazaki fragment is preceded by an RNA primer, which is displaced by the procession of the next Okazaki fragment during synthesis. In eukaryotic cells, a small amount of the DNA segment immediately upstream of the RNA primer is also displaced, creating a flap structure. This flap is then cleaved by endonucleases (such as Fen1, discussed later). At the replication fork, the gap in DNA after removal of the flap is sealed by DNA ligase I. Owing to the relatively short nature of the eukaryotic Okazaki fragment, DNA replication synthesis occurring discontinuously on the lagging strand is less efficient and more time consuming than leading-strand synthesis. Replication on the leading and lagging strands is performed by Pol ε and Pol δ, respectively. Many replisome factors (including the FPC [fork protection complex], Claspin, And1, and RFC [the replication factor C clamp loader]) are charged with regulating polymerase functions and coordinating DNA synthesis with the unwinding of the template strand by Cdc45-MCM [mini-chromosome maintenance]-GINS [go-ichi-ni-san]. The replisome also associates with checkpoint proteins as DNA replication and genome integrity surveillance mechanisms. Figure \(33\) shows an interactive iCn3D model of the Core human replisome (7PFO). (long load time) The leading DNA strand backbone is shown in spacefill magenta while the lagging strand backbone is shown in cyan. The DNA bases are shown as CPK spheres. The ATP analog, phosphaminophosphonic acid-adenlate ester, is shown in spacefill with CPK colors and labeled. The C-alpha traces of the different protein subunits are all shown in different colored alpha-C traces, except the DNA polymerase epsilon catalytic subunit A which is shown in cartoon form and colored by secondary structure. (long load time) At the eukaryotic replication fork, three distinct replicative polymerase complexes contribute to canonical DNA replication: α, δ, and ε. These three polymerases are essential for the viability of the cell. Because DNA polymerases require a primer on which to begin DNA synthesis, first, polymerase α (Pol α) acts as a replicative primase. Pol α is associated with an RNA primase and this complex accomplishes the priming task by synthesizing a primer that contains a short ~10-nucleotide RNA stretch followed by 10 to 20 DNA bases. Importantly, this priming action occurs at replication initiation at origins to begin leading-strand synthesis and also at the 5' end of each Okazaki fragment on the lagging strand. However, Pol α is not able to continue DNA replication. From in vitro studies, it was observed that DNA replication must be “handed off” to another polymerase to continue synthesis. The polymerase switching requires clamp loaders. Initially, it was thought that Pol δ performed leading-strand replication and that Pol α completed each Okazaki fragment on the lagging strand. Using mutator polymerase variants and mapping nucleotide misincorporation events, Kunkel and colleagues found that Pol ε and Pol δ mutations lead to mismatched nucleotide incorporation only on the leading and lagging strands, respectively. Thus, normal DNA replication requires the coordinated actions of three DNA polymerases: Pol α for priming synthesis, Pol ε for leading-strand replication, and Pol δ for generating Okazaki fragments during lagging-strand synthesis. In eukaryotes, DNA polymerases are grouped into seven families (A, B, C, D, X, Y, and RT). Crystal structures of the three nuclear replicative DNA polymerases demonstrate that they belong to the B family (Figure 25.1.17). All three replicative DNA polymerases are multi-subunit enzymes as shown in Table \(2\) below. Table 25.1.2 Subunits of the Major Eukaryotic Replicative DNA Polymerases Table \(2\): Subunits of the Major Eukaryotic Replicative DNA Polymerases. Doublié, S. and Zahn, K.E. (2014) Front. Microbiol 5:444 All B family polymerases are composed of five subdomains, the fingers, thumb, and palm which constitute the core of the enzyme, as well as an exonuclease domain and an N-terminal domain (NTD). The palm, a highly conserved fold composed of four antiparallel β strands and two helices, harbors two strictly conserved catalytic aspartates, located in motif A, DXXLYPS and motif C, DTDS , as shown in Figure \(34\). This fold is shared by a very large group of enzymes, including DNA and RNA polymerases, reverse transcriptases, CRISPR polymerase, and even reverse (3′–5′) transferases. In contrast, the thumb and the fingers subdomains exhibit substantially more structural diversity. The fingers undergo a conformational change upon binding DNA and the correct incoming nucleotide. This movement allows residues in the finger subdomain to come in contact with the nucleotide in the nascent base pair. The thumb holds the DNA duplex during replication and plays a part in processivity. The exonuclease domain carries a 3′–5′ proofreading activity, which removes misincorporated nucleotides. The NTD seems to be devoid of catalytic activity. In pol δ the NTD comprises three motifs, one has a topology resembling an OB fold, one a single-stranded DNA binding motif, and the another has a RNA-binding motif (RNA Recognition Motif or RRM). The NTD likely plays a role in polymerase stability and fidelity through its interactions with other domains. DNA polymerases require additional factors to support DNA replication in vivo. DNA polymerases have a semi-closed hand structure, which allows them to load onto DNA and translocate. This structure permits DNA polymerase to hold the single-stranded template, incorporate dNTPs at the active site, and release the newly formed double strand. However, the conformation of DNA polymerases does not allow for their stable interaction with the template DNA. To strengthen the interaction between template and polymerase, DNA sliding clamps have evolved, promoting the processivity of replicative polymerases. In eukaryotes, this sliding clamp is a homotrimer known as proliferating cell nuclear antigen (PCNA), which forms a ring structure. The PCNA ring has polarity with a surface that interacts with DNA polymerases and tethers them securely to DNA. PCNA-dependent stabilization of DNA polymerases has a significant effect on DNA replication because it enhances polymerase processivity up to 1,000-fold (Figure 25.1.19). The DNA helicases (MCM proteins) and polymerases must also remain in close contact at the replication fork (Figure 25.1.19). If unwinding occurs too far in advance of synthesis, large tracts of ssDNA are exposed. This can activate DNA damage signaling or induce aberrant DNA repair processes. To thwart these problems, the eukaryotic replisome contains specialized proteins that are designed to regulate the helicase activity ahead of the replication fork. These proteins also provide docking sites for physical interaction between helicases and polymerases, thereby ensuring that duplex unwinding is coupled with DNA synthesis. Control of Origin Firing Origin usage in eukaryotes can be dynamic, with origin firing at different sites depending on cell type and developmental stage. Nevertheless, the mechanism of replisome assembly and origin firing is highly conserved. During late mitosis and Gphase, cell cycle proteins, such as Cdc6, associate with Ori sites throughout the genome and recruit the helicase enzymes, MCMs 2-7 as shown in Figure \(35\). At this time, double hexamers of the MCM2-7 complex are loaded at replication origins. This generates a pre-replication complex (pre-RC). Origins with an associated pre-RC are considered licensed for replication. Licensed replication origins can then be “fired,” when replication actually initiates at the Ori. Origin firing is brought about by multiple phosphorylation events carried out by the cyclin E-CDK2 complex at the onset of S phase and by other cyclin-dependent kinases (CDKs) prior to individual origin firing (Figure \(35\)). Cyclin-dependent kinases (CDKs) are the families of protein kinases first discovered for their role in regulating the cell cycle. They are also involved in regulating transcription, mRNA processing, and the differentiation of nerve cells. CDKs are activated through the binding of an associated cyclin regulatory protein. Without a cyclin, CDKs exhibit little kinase activity. Following the phosphorylation of the pre-RC, origin melting occurs and DNA unwinding by the helicase generates ssDNA, exposing a template for replication (Figure \(35\)). The replisome then begins to form with the localization of replisome factors such as Cdc45. DNA synthesis begins on the melted template, and the replication machinery translocates away from the origin in a bidirectional manner. Pane (A) shows the combined activities of Cdc6 and Cdt1 bring MCM complexes (shown as blue circles of varying shades) to replication origins. Panel (B) shows CDK/DDK-dependent phosphorylation of pre-RC components leads to replisome assembly and origin firing. Cdc6 and Cdt1 are no longer required and are removed from the nucleus or degraded Panel (C) shows MCMs and associated proteins (GINS and Cdc45 are shown) unwinding DNA to expose template DNA. At this point, replisome assembly can be completed and replication initiated. “P” indicates phosphorylation. Replication through Nucleosomes Eukaryotic genomes are substantially more complicated than the smaller and unadorned prokaryotic genomes. Eukaryotic cells have multiple noncontiguous chromosomes, each of which must be compacted to allow packaging within the confined space of a nucleus. As seen in chapter 4, chromosomes are packaged by wrapping ~147 nucleotides (at intervals averaging 200 nucleotides) around an octamer of histone proteins, forming the nucleosome. The histone octamer includes two copies each of histone H2A, H2B, H3, and H4. In chapter 8, it was highlighted that histone proteins are subject to a variety of post-translational modifications, including phosphorylation, acetylation, methylation, and ubiquitination that represent vital epigenetic marks. The tight association of histone proteins with DNA in nucleosomes suggests that eukaryotic cells possess proteins that are designed to remodel histones ahead of the replication fork, in order to allow smooth progression of the replisome. It is also essential to reassemble histones behind the fork to reestablish the nucleosome conformation. Furthermore, it is important to transmit the epigenetic information found on the parental nucleosomes to the daughter nucleosomes, in order to preserve the same chromatin state. In other words, the same histone modifications should be present on the daughter nucleosomes as on the parental nucleosomes. This must all be done while doubling the amount of chromatin, which requires the incorporation of newly synthesized histone proteins. This process is accomplished by histone chaperones and histone remodelers, which are discussed below and shown in Figure \(36\). Histones are removed from chromatin ahead of the replication fork. FACT may facilitate this process. Asf1 recruits histone H3-H4 dimers to the replication fork. CAF-1 and Rtt106 load newly synthesized (light purple) histones to establish chromatin behind the fork. Previously loaded histones (dark purple) are also deposited on both daughter DNA strands. The histone chaperones involved in these processes are associated with replisome proteins: CAF-1/Rtt106 with PCNA and FACT/Asf1 with MCMs. Several histone chaperones are known to be involved in replication-coupled nucleosome assembly, including the FACT complex. The FACT complex components were originally identified as proteins that greatly stimulate transcription by RNA polymerase II. In budding yeast, FACT was found to interact with DNA Pol α-primase complex, and the FACT subunits were found to interact genetically with replication factors. More recently, studies showed that FACT facilitates DNA replication in vivo and is associated with the replisome in budding yeast and human cells. The FACT complex is a heterodimer that does not hydrolyze ATP, but facilitates the “loosening” of histones in nucleosomes Replication Fork Barriers and the Termination of Replication In prokaryotes, such as the E. coli, bidirectional replication initiates at a single replication origin on the circular chromosome and terminates at a site approximately opposed from the origin. This replication terminator region contains DNA sequences known as Ter sites, polar replication terminators that are bound by the Tus protein. The Ter-Tus complex counteracts helicase activity, resulting in replication termination. In this way, prokaryotic replication forks pause and terminate in a predictable manner during each round of DNA replication. In eukaryotes, the situation differs. Replication termination typically occurs by the collision of two replication forks anywhere between two active replication origins. The location of the collision can vary based on the replication rate of each of the forks and the timing of origin firing. Often, if a replication fork is stalled or collapsed at a specific site, replication of the site can be rescued when a replisome traveling in the opposite direction completes copying the region. However, there are numerous programmed replication fork barriers (RFBs) and replication “challenges” throughout the genome. To efficiently terminate or pause replication forks, some fork barriers are bound by RFB proteins in a manner analogous to E. coli Tus. In these circumstances, the replisome and the RFB proteins must specifically interact to stop replication fork progression. Telomeres and Replicative Senescence The End Replication Problem We have discussed the structure of telomers in the previous section. Let's look now at their activity/function. In humans, telomeres consist of hundreds to thousands of repetitive sequences of TTAGGG at chromosomal ends for maintaining genomic integrity. Because the DNA replication is asymmetric along double strands, RNA primer sequence at the 3′-hydroxyl end cannot be replaced by DNA polymerase I, as there is no 3'-OH primer group present for the polymerase to extend the DNA chain. This causes the loss of 30–200 nucleotides with each DNA replication and cell division and is known as the end replication problem. Telomeres provide a repetitive noncoding sequence of DNA at their 3′ en, to prevent the loss of critical genetically encoded information during replication. Moreover, telomeres are coated with a complex of six capping proteins, also known as shelterin proteins, which are packed into a compact T-loop structure that hides the ends of the chromosomes. This prevents the DNA repair machinery from mistaking chromosomal ends for double-stranded DNA breaks, as shown in Figure \(37\). Therefore, telomeres have been proposed as a mitotic clock that measures how many times a cell has divided and in essence, gives a cell a defined lifetime. Pane (A): shows telomeres located at the end of chromosomes, where they help protect against the loss of DNA during replication. Panel (B) shows DNA quadruplex formed by telomere repeats. The looped conformation of the DNA backbone is very different from the typical DNA helix, this is known as T-loop formation. The green spheres in the center represent potassium ions. The human telomerase enzyme is responsible for maintaining and elongating telomeres and consists of an RNA component (TERC) and a reverse transcriptase (TERT), that serves as the catalytic component, as shown in Figure \(38\). The TERT uses the TERC as a template to synthesize new telomeric DNA repeats at a single-stranded overhang to maintain telomere length (Figure 25.1.26). Some cells such as germ cells, stem cells, hematopoietic progenitor cells, activated lymphocytes, and most cancer cells constitutively express telomerase and maintain telomerase activity to overcome telomere shortening and cellular senescence. However, most other somatic cells generally have a low or undetectable level of telomerase activity and concomitantly limited longevity. Interestingly, overall telomerase activity decreases with age, but increases markedly in response to injury, suggesting a role for telomerase in cellular regeneration during wound healing. The telomere length and integrity are regulated through the interplay between the telomerase and shelterin proteins. The active site of the telomerase enzyme contains the RNA template, TERC (shown in red) and aligns with the last few telomeric bases at the end of the chromosome (shown in blue). This creates a single-stranded overhang that can be used as a template by the TERT reverse transcriptase to extend the telomere sequence. In vivo, shortened telomeres and damaged telomeres generally caused by reactive oxygen species (ROS) are usually assumed to be the main markers of cellular aging and are thought to be the main cause of replicative senescence. In vitro, telomeres lose approximately 50–200 bp at each division due to the end-replication problem. Approximately 100 mitoses are thought to be sufficient to reach the Hayflick limit, or the maximum number of mitotic events allowed prior to entering replicative senescence. Cells in continual renewal, such as blood cells, compensate for telomere erosion by expressing telomerase, the only enzyme able to polymerize telomeric sequences de novo at the extremity of telomeres. Knocking out telomerase components, such as the catalytic subunit (TERT) or the RNA template (TERC), induces several features of aging in mice. In humans, germline mutations in telomerase subunits are responsible for progeroïd syndromes, such as Dyskeratosis congenita, a rare genetic form of bone marrow failure. Furthermore, healthy lifespan in humans is positively correlated with longer telomere length and patients suffering from age-related diseases and premature aging have shorter telomeres compared with healthy individuals. An accumulation of unrepaired damage within telomeric regions has also been shown to accumulate in aging mice and non-human primates, suggesting that damage of telomeres with age may also be contributing to age-driven disease states and reduced health span. Thus, one could argue that the activation and expression of telomerase may be a way of reducing age-related diseases and increasing overall longevity. However, the constitutive expression of telomerase, unfortunately, is a characteristic of almost all cancer cells. It is therefore, no surprise that transgenic animals over-expressing the catalytic subunit of telomerase (mTERT), develop cancers earlier in life. However, over-expression of telomerase in mice that are highly resistant to cancers has shown large increases in median lifespan and significantly reduced age-associated disorders. Since humans are not highly resistant to cancer, this is not a feasible option for humans. However, additional studies in mice, where constitutive expression of telomerase is only introduced into a small percentage of host cells using adenovirus gene therapy techniques has yielded more promising results. Adenoviruses are a group of viruses that form an icosahedral protein capsid that houses a linear double-stranded DNA genome. Infections in humans typically cause symptoms of the common cold and are usually mild in nature. These are a good target for gene therapy, as the DNA that they carry can be mutated, so that they are deficient in their ability to replicate once they have infected the host. They can also be transformed to carry a gene of interest into the host, where that gene can then integrate into the host genome. Experiments in mice that were infected with an adenovirus carrying the mTERT gene showed that mTERT was delivered to a wide range of tissues within the body, and increased telomere length within those tissues. Furthermore, the mTERT expressing mice were healthier than their litter mates and displayed a reduction in disabling conditions associated with physiological aging such as osteoporosis and insulin resistance, as shown in Figure \(38\). Cognitive skills and metabolic functions were also improved. Noticeably, mice treated with gene therapy did not have increased incidence in cancer rates, suggesting that in at least for the short-lived mouse species, gene therapy approach to increased telomerase activity is safe. Within these animals, the median lifespan was increased by 24% when animals were treated at 1 year of age, and by 13% if treated at 2 years of age. Replication and Repair of Telomere Sequences In addition to the end replication problem, telomeric DNA (telDNA) replication and repair is a real challenge due to the different structural features of telomeres. First, the nucleotide sequence itself consists of a hexanucleotide motif (TTAGGG) repeated over kilobases, with the 5′-3′ strand named the “G-strand” due to its high content in guanine. During the progression of the replication fork, the lagging strand, corresponding to the G-strand, forms G-quadruplex (G4) structures, which have to be resolved to allow fork progression and to complete replication, as shown in Figure \(39\). Secondly, R-loops corresponding to highly stable RNA:DNA hybrids, involving the long non-coding telomeric transcript TERRA (telomeric repeat-containing RNA) also have to be dissociated. Thirdly, the extremity of telomeres adopts a specific loop structure, the T-loop, which has to be unraveled. This is the loop that hides the double-stranded end from the DNA damage sensors, and is locked by the hybridization of the 3′ single strand overhang extremity with the above 3′-5′ strand, thereby displacing the corresponding 5′-3′ strand to form a D-loop (displacement loop) structure (Figure \(38\)). Lastly, replication also has to deal with barriers encountered elsewhere in the genome, such as torsions and a condensed heterochromatic environment. Panel (a) shows the Telomeric sequence, with the G-strand in a solid red line and the C-strand in a solid green line, is depicted. The terminal D-loop structuring the much larger T-loop is stabilized by the shelterin complex. The replisome (PCNA, pol ε, etc) polymerizes a new G-strand (depicted in a dotted red line) and frees the parental G-strand, enabling the formation of G4 secondary structure. R-loops corresponding to TERRA hybridization (in dotted black lines) with the 3'-5' strand and torsions due to the fork progression are also shown. Panel (b) shows replication helpers, such as helicases, either helping in G4 unwinding or in D-loop unlocking are depicted. The DNAses (Top2a, DNA2) and RNAses (RNAse H1 and FEN1) help in resolving torsions and RNA:DNA heteroduplexes, while Timeless stimulates the replisome and POT1 competes with RPA1 for binding of the single-strand and helps in G4 dissolution. The shelterin components, POT1, TRF1, and TRF2 help in loading the helper proteins (fine green arrows) Since telomeres face a host of obstacles to completing the replication process, as discussed in Figure 25.1.28, the cell possesses a set of specialized machinery to fully achieve their replication, such as the RTEL1, TRF1, and TRF2 proteins, DNAses, RNAsses, and Timeless. The recruitment of these factors is orchestrated by the shelterin complex. At the molecular level, the GGG telomeric repeats are particularly sensitive to ROS, which produce stretches of 8-oxoguanine that are especially difficult to repair. Coupled with inefficient telomere repair, these ROS-induced lesions produce single and double-strand breaks, and/or generate replicative stress, ultimately resulting in telomere shortening. The presence of unrepaired single or tandem 8-oxoguanine drastically inhibits the binding of TRF1 and TRF2, and impairs the recruitment of telomerase, especially when ROS damage is localized in the 3′ overhang. This type of damage contributes to telomere deprotection and shortening. Strikingly, ROS (and other metabolic stresses) also induce the relocation of TERT to mitochondria, as observed (i) in primary neurons after oxidative stress; (ii) in neurons exposed to the tau protein; (iii) in Purkinje neurons subjected to excitotoxicity; and (iv) in cancer cell lines treated with a G4 ligand. Mitochondrial TERT increases the inner membrane potential, as well as the mtDNA copy number, and decreases ROS production with a protective effect on mtDNA. Mitochondria are also critical sensors of cellular damage and contribute to the processes of autophagy and apoptosis (programmed cell death). The relocalization of TERT following chromosomal damage in the nucleus, may indicate one mechanism the mitochondria utilizes to monitor cellular stress and damage. Replication of Mitochondrial DNA Mammalian mitochondria contain multiple copies of a circular, double-stranded DNA genome approximately 16.6 kb in length, as shown in Figure \(40\). The two strands of mtDNA differ in their base composition, with one being rich in guanines, making it possible to separate a heavy (H) and a light (L) strand by density centrifugation. The mtDNA contains one longer noncoding region (NCR) also referred to as the control region. In the NCR, there are promoters for polycistronic transcription, one for each mtDNA strand; the light strand promoter (LSP) and the heavy strand promoter (HSP). The NCR also harbors the origin for H-strand DNA replication (OH). A second origin for L-strand DNA replication (OL) is located outside the NCR, within a tRNA cluster. Falkenberg, M. (2018) Essays Biochem 62(3):287-296 As shown in Figure \(40\), the genome encodes for 13 mRNA (green), 22 tRNA (violet), and 2 rRNA (pale blue) molecules. There is also a major noncoding region (NCR), which is shown enlarged at the top in blue. The major NCR contains the heavy strand promoter (HSP), the light strand promoter (LSP), three conserved sequence boxes (CSB1-3, orange), the H-strand origin of replication (OH), and the termination-associated sequence (TAS, yellow). The triple-stranded displacement-loop (D-loop) structure is formed by a premature termination of nascent H-strand DNA synthesis at TAS. The short H-strand replication product formed in this manner is termed 7S DNA. A minor NCR, located approximately 11,000 bp downstream of OH, contains the L-strand origin of replication (OL). A dedicated DNA replication machinery is required for its maintenance. Mammalian mtDNA is replicated by proteins distinct from those used for nuclear DNA replication and many are related to replication factors identified in bacteriophages. DNA polymerase γ (POLγ) is the replicative polymerase in mitochondria. In human cells, POLγ is a heterotrimer with one catalytic subunit (POLγA) and two accessory subunits (POLγB). POLγA belongs to the A family of DNA polymerases and contains a 3′–5′ exonuclease domain that acts to proofread the newly synthesized DNA strand. POLγ is a highly accurate DNA polymerase with a frequency of misincorporation lower than 1 × 10−6. The accessory POLγB subunit enhances interactions with the DNA template and increases both the catalytic activity and the processivity of POLγA. The DNA helicase TWINKLE travels in front of POLγ, unwinding the double-stranded DNA template. TWINKLE forms a hexamer and requires a fork structure (a single-stranded 5′-DNA loading site and a short 3′-tail) to load and initiate unwinding. Mitochondrial single-stranded DNA-binding protein (mtSSB) binds to the formed ssDNA, protects it against nucleases, and prevents secondary structure formation The most accepted model of DNA replication in the mitochondria is the strand displacement model, as shown in Figure \(41\). Within this model, DNA synthesis is continuous on both the H- and L-strand. There is a dedicated origin for each strand, OH and OL. First, replication is initiated at OH and DNA synthesis then proceeds to produce a new H-strand. During the initial phase, there is no simultaneous L-strand synthesis and mtSSB covers the displaced, parental H-strand. By binding to single-stranded DNA, mtSSB prevents the mitochondrial RNA polymerase (POLRMT) from initiating random RNA synthesis on the displaced strand. When the replication fork has progressed about two-thirds of the genome, it passes the second origin of replication, OL. When exposed in its single-stranded conformation, the parental H-strand at OL folds into a stem–loop structure. The stem efficiently blocks mtSSB from binding and a short stretch of single-stranded DNA in the loop region, therefore remains accessible, allowing POLRMT to initiate RNA synthesis. POLRMT is not processive on single-stranded DNA templates. After adding approximately 25 nucleotides, it is replaced by POLγ and L-strand DNA synthesis is initiated. From this point, H- and L-strand synthesis proceeds continuously until the two strands have reached full circle. Replication of the two strands is linked, since H-strand synthesis is required for the initiation of L-strand synthesis. DNA Ligase III is used to complete the ligation of the newly formed DNA strands. During DNA replication, the parental molecule remains intact, which poses a steric problem for the moving replication machinery. Topoisomerases belonging to the type 1 family can relieve torsional strain formed in this way, by allowing one of the strands to pass through the other. In mammalian mitochondria, TOP1MT a type IB enzyme can act as a DNA “swivel”, working together with the mitochondrial replisome. Furthermore, replication of circular DNA often causes the formation of catenanes, or interlocked circles that need to be separated from one another. The type 1A topoisomerase, topoisomerase 3α (Top3α), is required to resolve the hemicatenane structure that can form during mtDNA replication. Curiously, not all replication events initiated at OH continue to full circle. Instead, 95% are terminated after about the first 650 nucleotides at a sequence known as the termination associated sequences (TAS) (Figure 25.1.23). This creates a short DNA fragment known as the 7S DNA, that remains bound to the parental L-strand, while the parental H-strand is displaced (Figure 25.1.23). As a result, a triple-stranded displacement loop structure, a D-loop, is formed. The functional importance of the D-loop structure is unclear and how replication is terminated at TAS is also not known. Mastering the Content Which of the following is the enzyme that replaces the RNA nucleotides in a primer with DNA nucleotides? 1. DNA polymerase III 2. DNA polymerase I 3. primase 4. helicase [reveal-answer q=”628075″]Show Answer[/reveal-answer] [hidden-answer a=”628075″]Answer b. DNA polymerase I is the enzyme that replaces the RNA nucleotides in a primer with DNA nucleotides.[/hidden-answer] Which of the following is not involved in the initiation of replication? 1. ligase 2. DNA gyrase 3. single-stranded binding protein 4. primase [reveal-answer q=”820951″]Show Answer[/reveal-answer] [hidden-answer a=”820951″]Answer a. Ligase is not involved in the initiation of replication.[/hidden-answer] Which of the following enzymes involved in DNA replication is unique to eukaryotes? 1. helicase 2. DNA polymerase 3. ligase 4. telomerase [reveal-answer q=”650146″]Show Answer[/reveal-answer] [hidden-answer a=”650146″]Answer d. Telomerase is unique to eukaryotes.[/hidden-answer] Which of the following would be synthesized using 5′-CAGTTCGGA-3′ as a template? 1. 3′-AGGCTTGAC-4′ 2. 3′-TCCGAACTG-5′ 3. 3′-GTCAAGCCT-5′ 4. 3′-CAGTTCGGA-5′ [reveal-answer q=”429167″]Show Answer[/reveal-answer] [hidden-answer a=”429167″]Answer c. 3′-GTCAAGCCT-5′[/hidden-answer] The enzyme responsible for relaxing supercoiled DNA to allow for the initiation of replication is called ________. [reveal-answer q=”855893″]Show Answer[/reveal-answer] [hidden-answer a=”855893″]The enzyme responsible for relaxing supercoiled DNA to allow for the initiation of replication is called DNA gyrase or topoisomerase II.[/hidden-answer] Unidirectional replication of a circular DNA molecule like a plasmid that involves nicking one DNA strand and displacing it while synthesizing a new strand is called ________. [reveal-answer q=”378861″]Show Answer[/reveal-answer] [hidden-answer a=”378861″]Unidirectional replication of a circular DNA molecule like a plasmid that involves nicking one DNA strand and displacing it while synthesizing a new strand is calledrolling circle replication.[/hidden-answer] More primers are used in lagging strand synthesis than in leading strand synthesis. [reveal-answer q=”25479″]Show Answer[/reveal-answer] [hidden-answer a=”25479″]True[/hidden-answer] 1. Why is primase required for DNA replication? 2. What is the role of single-stranded binding protein in DNA replication? 3. Below is a DNA sequence. Envision that this is a section of a DNA molecule that has separated in preparation for replication, so you are only seeing one DNA strand. Construct the complementary DNA sequence (indicating 5′ and 3′ ends).DNA sequence: 3′-T A C T G A C T G A C G A T C-5′ 4. Review Figure 1 and Figure 2. Why was it important that Meselson and Stahl continue their experiment to at least two rounds of replication after isotopic labeling of the starting DNA with15N, instead of stopping the experiment after only one round of replication? 5. If deoxyribonucleotides that lack the 3′-OH groups are added during the replication process, what do you expect will occur?

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/24%3A_DNA_Metabolism/24.01%3A_DNA_Replication.txt

Search Fundamentals of Biochemistry The integrity of the DNA structure for cell viability is underscored by the vast amounts of cellular machinery dedicated to ensuring its accurate replication, repair, and storage. Even still, mutations within the DNA are a fairly common event. DNA Mutations Mutations are random changes that occur within the sequence of bases in DNA. They can be large-scale, altering the structure of the chromosomes, or small scale where they only alter a few or even a single base or nucleotide. Mutations can occur for many reasons. For example, DNA mutations can be caused by mistakes made by the DNA polymerase during replication. As noted in chapter 9, DNA polymerases are highly processive enzymes that contain proofreading and editing functions. With these safeguards, their error rates are typically very low and range from one in a million bases to one in a billion bases. Even with such high fidelity, this error rate will lead to between 3 and 3,000 errors within the human genome for each cell undergoing DNA replication. DNA mutations can also result through the replication of DNA that has been damaged by endogenous or exogenous agents. The next section will highlight common types of DNA damage and their effects. If a DNA polymerase encounters a damaged DNA base in the template DNA during replication it may place a random nucleotide base across from the lesion. For example, an adenine-containing nucleotide will often be added across a lesion, regardless of what the correct match should be. This can lead to the formation of transition or transversion mutations. A transition mutation is a point mutation that changes a purine nucleotide to another purine (A ↔ G) or a pyrimidine nucleotide to another pyrimidine (C ↔ T). Transversion refers to the substitution of a purine for a pyrimidine or vice versa. Sometimes lesions may cause bases to be skipped during replication or cause extra nucleotides to be inserted into the backbone. DNA polymerases can also slip during the replication of regions of the DNA that have repeated sequences or large stretches repeating a single base. Larger lesions or cross-links in the DNA during replication can lead to more catastrophic DNA damage including DNA strand breaks. Mutations may also occur during the processes of mitosis and meiosis when sister chromatids and/or homologous chromosomes are separated from one another. In nature, mutagenesis, or the process of generating DNA mutations, can lead to changes that are harmful, beneficial, or have no effect. Harmful mutations can lead to cancer and various heritable diseases, but beneficial mutations are the driving force of evolution. In 1927, Hermann Muller first demonstrated the effects of mutations with observable changes in chromosomes. He induced mutagenesis by irradiating fruit flies with X-rays. When a mutation is caused by an environmental factor or a chemical agent, that agent is called a mutagen. Typical mutagens include chemicals, like those inhaled while smoking, and radiation, such as X-rays, ultraviolet light, and nuclear radiation. Different mutagens have different modes of damaging DNA and are discussed further in the next section. It is important to note that DNA damage, in and of itself, does not necessarily lead to the formation of a mutation in the DNA. There are elaborate DNA repair processes designed to recognize and repair different types of DNA lesions. Fewer than 1 in 1,000 DNA lesions will result in a DNA mutation. The processes of DNA damage recognition and repair are the focus of later sections within this chapter. Types of Mutations There are a variety of types of mutations. Two major categories of mutations are germline mutations and somatic mutations. • Germline mutations occur in gametes, the sex cells, such as eggs and sperm. These mutations are especially significant because they can be transmitted to offspring and every cell in the offspring will have the mutations. • Somatic mutations occur in other cells of the body. These mutations may have little effect on the organism because they are confined to just one cell and its daughter cells. Somatic mutations also cannot be passed on to offspring. Mutations also differ in the way that the genetic material is changed. Mutations may change an entire chromosome or just one or a few nucleotides. Chromosomal alterations are mutations that change chromosome structure or number. They occur when a section of a chromosome breaks off and rejoins incorrectly or does not rejoin at all. Possible ways these mutations can occur are illustrated in the figure below. Chromosomal alterations are very serious. They often result in the death of the cell or organism in which they occur. If the organism survives, it may be affected in multiple ways. An example of a human chromosomal alteration is the mutation that causes Down Syndrome. It is a duplication mutation that leads to developmental delays and other abnormalities. It occurs when the individual inherits an extra copy of chromosome 21. It is also called trisomy ("three-chromosome") 21. Thus, large-scale mutations in the chromosomal structure include (1) Amplifications (including gene duplications) where repetition of a chromosomal segment or presence of an extra piece of a chromosome broken piece of a chromosome may become attached to a homologous or non-homologous chromosome so that some of the genes are present in more than two doses leading to multiple copies of all chromosomal regions, increasing the dosage of the genes located within them, (2) Deletions of large chromosomal regions, leading to loss of the genes within those regions, and (3) Chromosomal Rearrangements such as translocations (which interchange of genetic parts from nonhomologous chromosomes), insertions (which insert segments of one chromosome into another nonhomologous chromosome), and inversions (which invert or flip a section of a chromosome into the opposite orientation), as shown in Figure \(1\). There are also smaller mutations that can occur that only alter a single nucleotide or a small number of nucleotides within a localized region of the DNA. These are classified according to how the DNA molecule is altered. One type, a point mutation, affects a single base and most commonly occurs when one base is substituted or replaced by another. Mutations also result from the addition of one or more bases, known as an insertion, or the removal of one or more bases, known as a deletion. Point mutations (Table \(1\) and Figure \(2\)) may have a wide range of effects on protein function. As a consequence of the degeneracy of the genetic code, a point mutation will commonly result in the same amino acid being incorporated into the resulting polypeptide despite the sequence change. This change would not affect the protein’s structure and is thus called a silent mutation. A missense mutation results in a different amino acid being incorporated into the resulting polypeptide. The effect of a missense mutation depends on how chemically different the new amino acid is from the wild-type amino acid. The location of the changed amino acid within the protein also is important. For example, if the changed amino acid is part of the enzyme’s active site or greatly affects the shape of the enzyme, then the effect of the missense mutation may be significant. Many missense mutations result in proteins that are still functional, at least to some degree. Sometimes the effects of missense mutations may be only apparent under certain environmental conditions; such missense mutations are called conditional mutations. Rarely, a missense mutation may be beneficial. Under the right environmental conditions, this type of mutation may give the organism that harbors it a selective advantage. Yet another type of point mutation called a nonsense mutation, converts a codon encoding an amino acid (a sense codon) into a stop codon (a nonsense codon). Nonsense mutations result in the synthesis of proteins that are shorter than the wild type and typically not functional. Table \(1\): Types of Point Mutations Type Description Example Effect Silent mutated codon codes for the same amino acid CAA (glutamine) → CAG (glutamine) none Missense mutated codon codes for a different amino acid CAA (glutamine) → CCA (proline) variable Nonsense a mutated codon is a premature stop codon CAA (glutamine) → UAA (stop) usually serious Smaller scale deletions and insertions also cause various effects. Because codons are triplets of nucleotides, insertions or deletions in groups of three nucleotides may lead to the insertion or deletion of one or more amino acids and may not cause significant effects on the resulting protein’s functionality. However, frameshift mutations, caused by insertions or deletions of a number of nucleotides that are not a multiple of three are extremely problematic because a shift in the reading frame results, as shown in Figure \(3\). Because ribosomes read the mRNA in triplet codons, frameshift mutations can change every amino acid after the point of the mutation. The new reading frame may also include a stop codon before the end of the coding sequence. Consequently, proteins made from genes containing frameshift mutations are nearly always nonfunctional. The majority of mutations have neither negative nor positive effects on the organism in which they occur. These mutations are called neutral mutations. Examples include silent point mutations, which are neutral because they do not change the amino acids in the proteins they encode. Some mutations have a positive effect on the organism in which they occur. They are referred to as beneficial mutations. If they occur in germline cells (eggs or sperm) these traits can be heritable and passed from one generation to the next. Beneficial mutations generally code for new versions of proteins that help organisms adapt to their environment. If they increase an organism’s chances of surviving or reproducing, the mutations are likely to become more common within a population over time. There are several well-known examples of beneficial mutations. Here are just two: 1. Mutations have occurred in bacteria that allow the bacteria to survive in the presence of antibiotic drugs. The mutations have led to the evolution of antibiotic-resistant strains of bacteria. 2. A unique mutation is found in people in a small town in Italy. The mutation protects them from developing atherosclerosis, which is the dangerous buildup of fatty materials in blood vessels. The individual in which the mutation first appeared has even been identified. Harmful mutations can also occur. Imagine making a random change in a complicated machine such as a car engine. The chance that the random change would improve the functioning of the car is very small. The change is far more likely to result in a car that does not run well or perhaps does not run at all. By the same token, any random change in a gene's DNA is more likely to result in the production of a protein that does not function normally or may not function at all, than in a mutation that improves the function. Such mutations are likely to be harmful. Harmful mutations may cause genetic disorders or cancer. • A genetic disorder is a disease, syndrome, or other abnormal condition caused by a mutation in one or more genes or by a chromosomal alteration. An example of a genetic disorder is cystic fibrosis. A mutation in a single gene causes the body to produce thick, sticky mucus that clogs the lungs and blocks ducts in digestive organs. Genetic disorders are usually caused by gene mutations that occur within germline cells and are heritable. • Illnesses caused by mutations that occur within an individual, but are not passed on to their offspring, are mutations that occur in somatic cells. Cancer is a disease caused by an accumulation of mutations within somatic cells. It results in cells that grow out of control and form abnormal masses of cells called tumors. It is generally caused by mutations in genes that regulate the cell cycle, DNA repair, angiogenesis, and other genes that favor cell growth and survival. Because of the mutations, cells with the mutated DNA have evolved to divide without restrictions, hide from the immune system, and develop drug resistance. Types of DNA Damage DNA damage, due to environmental factors and normal metabolic processes inside the cell, occurs at a rate of 1,000 to 1,000,000 molecular lesions per cell per day. While this constitutes only 0.000165% of the human genome's approximately 6 billion bases (3 billion base pairs), if left unrepaired can cause mutations in critical genes (such as tumor suppressor genes) can impede a cell's ability to carry out their function and appreciably increase the likelihood of tumor formation and disease states such as cancer. The vast majority of DNA damage affects the primary structure of the double helix; that is, the bases themselves are chemically modified. These modifications can, in turn, disrupt the molecules' regular helical structure by introducing non-native chemical bonds or bulky adducts that do not fit in the standard double helix. Unlike proteins and RNA, DNA usually lacks tertiary structure, and therefore damage or disturbance does not occur at that level. DNA is, however, supercoiled and wound around "packaging" proteins called histones (in eukaryotes), and both superstructures are vulnerable to the effects of DNA damage. Several types of DNA damage can occur due either to normal cellular processes or due to the environmental exposure of cells to DNA-damaging agents. DNA bases can be damaged by: (1) oxidative processes, (2) alkylation of bases, (3) base loss caused by the hydrolysis of bases, (4) bulky adduct formation, (5) DNA crosslinking, and (6) DNA strand breaks, including single and double-stranded breaks. An overview of these types of damage is described below. Oxidative Damage Reactive oxygen species (ROS) can cause significant cellular stress and damage including oxidative DNA damage. Hydroxyl radicals (OH) are one of the most reactive and electrophilic of the ROS and can be produced by ultraviolet and ionizing radiations or from other radicals arising from enzymatic reactions. The OH can cause the formation of 8-oxo-7,8-dihydroguanine (8-oxoG) from guanine residues, among other oxidative products, as shown in Figure \(4\). Guanine is the most easily oxidized of the nucleic acid bases because it has the lowest ionization potential among the DNA bases. The 8-oxo-dG is one of the most abundant DNA lesions, and it is considered as a biomarker of oxidative stress. It has been estimated that up to 100,000 8-oxo-dG lesions can occur daily in DNA per cell. The reduction potential of 8-oxo-dG is even lower (0.74 V vs. NHE) than that of guanosine (1.29 V vs NHE). Therefore, it can be further oxidized creating a variety of secondary oxidation products. As mentioned previously, increased levels of 8-oxo-dG in tissue can serve as a biomarker of oxidative stress. Furthermore, increased levels of 8-oxo-dG are frequently found associated with carcinogenesis and other disease states, as shown in Figure \(5\). During the replication of DNA that contains 8-oxo-dG, adenine is most often incorporated across from the lesion. Following replication, the 8-oxo-dG is excised during the repair process and thymine is incorporated in its place. Thus, 8-oxo-dG mutations typically result in a G to T transversion. Alkylation of Bases Alkylating agents are widespread in the environment and are also produced endogenously, as by-products of cellular metabolism. They introduce lesions into DNA or RNA bases that can be cytotoxic, mutagenic, or neutral to the cell. Figure \(6\) depicts the major reactive sites on the DNA bases that are susceptible to alkylation. Cytotoxic lesions block replication, interrupt transcription, or signal the activation of apoptosis, whereas mutagenic ones are miscoding and cause mutations in newly synthesized DNA. The most common type of alkylation is methylation with the major products including N7-methylguanine (7meG), N3-methyladenine (3meA), and O6-methylguanine (O6meG). Smaller amounts of methylation also occurs on other DNA bases, and include the formation of N1-methyladenine (1meA), N3-methylcytosine (3meC), O4-methylthymine (O4meT), and methyl phosphotriesters (MPT). Alkylating agents can cause damage to all exocyclic nitrogens and oxygens in DNA and RNA, as well as at ring nitrogens (Figure 25.2.6A). However, the percentage of each base site modified depends on the alkylating agent, the position in DNA or RNA, and whether nucleic acids are single- or double-stranded. Interestingly, O-alkylations are more mutagenic and harmful than N-alkylations, which may be more cytotoxic, but not as mutagenic. As we will explore in Chapter 13, methylation of DNA also serves as an important mechanism regulating gene expression. Base Loss An AP site (apurinic/apyrimidinic site), also known as an abasic site, is a location in DNA (also in RNA but much less likely) that has neither a purine nor a pyrimidine base, either spontaneously or due to DNA damage, as shown in Figure \(7\). It has been estimated that under physiological conditions 10,000 apurinic sites and 500 apyrimidinic may be generated in a cell daily. AP sites can be formed by spontaneous depurination, but also occur as intermediates in base excision repair, the repair process described in section 25.2.5. If left unrepaired, AP sites can lead to mutation during semiconservative replication. They can cause replication fork stalling and are often bypassed by translesion synthesis, which is discussed in greater detail in section 12.8. In E. coli, adenine is preferentially inserted across from AP sites, known as the "A rule". The situation is more complex in higher eukaryotes, with different nucleotides showing a preference depending on the organism and environmental conditions. Bulky Adduct Formation Some chemicals are biologically reactive and will form covalent linkages with biological molecules such as DNA and proteins creating large bulky adducts, or appendages, that branch off from the main molecule. We will use the mutagen/carcinogen, benzo[a]pyrene, as an example for this process. Benzo[a]pyrene is a polycyclic aromatic hydrocarbon that forms during the incomplete combustion of organic matter at temperatures between 300°C (572°F) and 600°C (1,112°F). The ubiquitous compound can be found in coal tar, tobacco smoke, and many foods, especially grilled meats. Benzo[a]pyrene is a procarcinogen that needs to be biologically activated by metabolism before it forms a reactive metabolite, as in Figure \(8\). Normally, when the body is exposed to foreign molecules, it will start a metabolic process that makes the molecule more hydrophilic and easier to remove as a waste product. Unfortunately, in the case of benzo[a]pyrene, the resulting metabolite is a highly reactive epoxide that forms a bulky adduct preferentially with guanine residues in DNA. If left unrepaired, during DNA replication an adenine will usually be placed across from the lesion in the daughter molecule. Subsequent repair of the adduct will result in the replacement of the damaged guanine base with thymine, causing a G --> T transversion mutation. DNA Crosslinking Crosslinking of DNA occurs when various exogenous or endogenous agents react with two nucleotides of DNA, forming a covalent linkage between them. This crosslink can occur within the same strand (intrastrand) or between opposite strands of double-stranded DNA (interstrand), as shown in Figure \(9\). These adducts interfere with cellular metabolism, such as DNA replication and transcription, triggering cell death. UV light can cause molecular crosslinks to form between two pyrimidine residues, commonly two thymine residues, that are positioned consecutively within a strand of DNA, as shown in Figure \(10\). Two common UV products are cyclobutane pyrimidine dimers (CPDs) and 6–4 photoproducts. These premutagenic lesions alter the structure and possibly the base pairing. Up to 50–100 such reactions per second might occur in a skin cell during exposure to sunlight, but are usually corrected within seconds by photolyase reactivation or nucleotide excision repair. Uncorrected lesions can inhibit polymerases, cause misreading during transcription or replication, or lead to the arrest of replication. Pyrimidine dimers are the primary cause of melanomas in humans. DNA Strand Breaks Ionizing radiation such as that created by radioactive decay or in cosmic rays causes breaks in DNA strands (see Figure above). Low-level ionizing radiation may induce irreparable DNA damage (leading to replication and transcription errors needed for neoplasia or may trigger viral interactions) leading to premature aging and cancer. Chemical agents that form crosslinks within the DNA, especially interstrand crosslinks, can also lead to DNA strand breaks if the damaged DNA undergoes DNA replication. Crosslinked DNA can cause topoisomerase enzymes to stall in the transition state when the DNA backbone is cleaved. Instead of relieving supercoiling and resealing the backbone, the stalled topoisomerase remains covalently linked to the DNA in a process called abortive catalysis. This leads to the formation of a single-stranded break in the case of Top1 enzymes or double-stranded breaks in the case of Top2 enzymes. DNA double-strand breaks due to topoisomerase stalling can also occur during the transcription of DNA, as shown in Figure \(11\). Abortive catalysis and the formation of DNA strand breaks during transcriptional events may serve as a damage sensor within the cell and help to instigate DNA damage response signaling pathways that initiate DNA repair processes. Panel (1) shows that in the uninduced state of transcription, Pol II is paused between +25 and +100 from the transcription start site. The pausing is attributed to different elements including pausing-stabilizing transcription factors, the +1 nucleosome, and DNA structure and torsion. Positive supercoiling ahead of Pol II may require the function of TOP2B. Panel (2) shows transcription activation induced by various stimuli activates TOP2B to resolve DNA torsion in the promoter and gene body. Panel (3) shows that in this process, double-strand breaks could be formed from abortive catalysis of TOP2B, which occurs frequently in some genes. This may be responsible for DNA damage response signaling that has been observed in a number of stimulus-inducible genes in humans. Figure from: Cellular Stress and DNA Damage Response Genetic damage produced by either exogenous or endogenous mechanisms represents an ongoing threat to the cell. To preserve genome integrity, eukaryotic cells have evolved repair mechanisms specific for different types of DNA Damage. However, regardless of the type of damage a sophisticated surveillance mechanism, that elicits DNA damage checkpoints, detects and signals its presence to the DNA repair machinery. DNA damage checkpoints have been functionally conserved throughout eukaryotic evolution, with most of the relevant players in the checkpoint response highly conserved from yeast to humans. Checkpoints are induced to delay cell cycle progression and to allow cells time to repair damaged DNA before DNA replication, as shown in Figure \(12\). Once the damaged DNA is repaired, the checkpoint machinery triggers signals that will resume cell cycle progression. Within cells, multiple pathways contribute to DNA repair, but independent of the specific repair pathway involved, three phases of checkpoint activation are traditionally identified: (1) Sensing of damage, (2) Activating the signaling cascade, and (3) Switching on downstream effectors. The sensor phase recognizes the damage and activates the signal transduction phase to block cell cycle progression and select the appropriate repair pathway. In addition to blocking cell cycle progression, DNA damage sensors also activate DNA repair mechanisms that are specific for the type of damage present. For example, single-stranded DNA breaks are repaired primarily by Base Excision Repair, bulky DNA adducts, and crosslinks are repaired by Nucleotide Excision Repair, and smaller nucleotide mutations, such as alkylation are repaired by Mismatch Repair. Cells also have two major mechanisms for repairing Double-Strand-Breaks (DSBs). They include Non-Homologous End-Joining (NHEJ) and Homologous Recombination (HR). If damage is too extensive to be repaired, apoptotic pathways will be elicited. In the following sections, details about the major DNA repair pathways will be given. In multicellular organisms, the response to DNA damage can result in two major physiological consequences: (1) Cells can undergo cell cycle arrest, repair the damage, and re-enter the cell cycle, or (2) cells can be targeted for cell death (apoptosis) and removed from the population. The cell cycle process is highly conserved and precisely controlled to govern the genome duplication and separation into the daughter cells. The cell cycle consists of four distinct and ordered phases, termed G0/G1 (gap 1), S (DNA synthesis), G2 (gap 2), and M (mitosis). Multiple checkpoints exist within each stage of the cell cycle to ensure the faithful replication of DNA in the S phase and the precise separation of the chromosomes into daughter cells. The G1 and G2 phases are critical regulatory checkpoints, whereby the restriction point between the G1 and S phase determines whether the cells enter the S phase or exit the cell cycle to halt at the G0 phase. The cell cycle progression requires the activity of cyclin-dependent kinases (CDKs), a group of serine/threonine kinases. CDKs are activated when they form complexes with cyclin regulatory proteins that are expressed specifically at different stages of the cell cycle. Cyclins bind to and stabilize CDKs in their active conformation. The formation of cyclin/CDKs controls the cell-cycle progression via phosphorylation of the target genes, such as tumor suppressor protein retinoblastoma (Rb). During DNA damage, the cell cycle is arrested or blocked by the action of cyclin-dependent kinase inhibitors. As noted in Figure 12.12, this is a complicated signal transduction cascade that has many downstream effects. A primary function of cell cycle arrest is that CDK inhibition allows time for DNA repair before cell-cycle progression into the S-phase or mitosis. As shown in Figure 25.2.12, two major cell-cycle checkpoints respond to DNA damage; they occur pre- and post-DNA synthesis in the G1 and G2 phases, respectively, and impinge on the activity of specific CDK complexes. The checkpoint kinases phosphatidylinositol 3-kinase (PI3K)-like protein kinases (PI3KKs) ataxia telangiectasia and Rad3-related (ATR) or ataxia telangiectasia mutated (ATM) protein, and the transducer checkpoint kinases CHK1 (encoded by the CHEK1 gene) and CHK2 (encoded by the CHEK2 gene) are key regulators of DNA damage signaling. The DNA damage signaling is detected by ATM/ATR, which then phosphorylates and activates CHK2/CHK1, respectively. The activated CHK2 is involved in the activation of p53, leading to p53-dependent early phase G1 arrest to allow time for DNA repair. The activation of p53 induces the expression of the Cyclin-Dependent Kinase Inhibitor (CKI) p21CIP1 gene, leading to the inhibition of cyclin E/CDK2 complexes and subsequent upregulation of DNA repair machinery. If the DNA repair cannot be completed successfully or the cells cannot program to respond to the stresses of viable cell-cycle arrest, the cells face the fate of apoptosis induced by p53. The activated CHK1 mediates temporary S phase arrest through phosphorylation to inactivate CDC25A, causing ubiquitination and proteolysis. Moreover, the activated CHK1 phosphorylates and inactivates CDC25C, leading to cell-cycle arrest in the G2 phase. The active CHK1 also directly stimulates the phosphorylation of WEE1, resulting in enhancing the inhibitory Tyr15 phosphorylation of CDK2 and CDK1 and subsequent cell-cycle blocking in the G2 phase. The activity of WEE1 can also be stimulated by the low levels of CDK activity in the G2 cell-cycle phase. The SAC, also known as the mitotic checkpoint, functions as the monitor of the correct attachment of the chromosomes to the mitotic spindle in metaphase, which is regulated by the TTK protein kinase (TTK, also known as monopolar spindle 1 (MPS1)). The activation of SAC transiently induces cell-cycle arrest by inhibiting the activation of APC/C. To establish and maintain the mitotic checkpoint, the TTK recruits many checkpoint proteins to kinetochores during mitosis via phosphorylating its substrates to ensure adequate chromosome segregation and genomic integrity. In this way, the genomic instability from chromosome segregation defects is protected by SAC. Once the SAC is passed, the APC/C E3 ligase complex stimulates and tags cyclin B and securin for ubiquitin-mediated degradation, leading to the initiation of mitosis. In a word, the checkpoints offer a failsafe mechanism to ensure the genomic integrity from the parental cell to the daughter cell. The signal transduction cascade of checkpoint activation eventually converges to CDK inhibition, which indicates the CDK function as a key driver of cell-cycle progression. Mismatch Repair DNA mismatch repair (MMR) is a highly conserved DNA repair system that greatly contributes to maintaining genome stability through the correction of mismatched base pairs and small modifications of DNA bases, such as alkylation. The major source of mismatched base pairs is replication error, although it can arise also from other biological processes. Thus, the MMR machinery must have a mechanism for determining which strand of the DNA is the template strand and which strand has been newly synthesized. In E. coli, methylation of the DNA is a common post-replicative modification that occurs. Thus, in newly synthesized DNA, the unmethylated strand is recognized as the new strand, and the methylated strand is used as the template to repair mismatches. In E.coli, MMR increases the accuracy of DNA replication by 20–400-fold. Mutations and epigenetic silencing in MMR genes have been implicated in up to 90% of human hereditary nonpolyposis colon cancers, indicating the significance of this repair system in maintaining genomic stability. Post-replicative MMR is performed by the long-patch MMR mechanism in which multiple proteins are involved and a relatively long tract of the oligonucleotide is excised during the repair reaction. In contrast, particular kinds of mismatched base pairs are repaired through very short-patch MMR in which a short oligonucleotide tract is excised to remove the lesion. Table\(2\) below shows mismatch repair enzymes in bacteria, yeast, and humans. MMR in eukaryotes and most bacteria directs the repair to the error-containing strand of the mismatched duplex by recognizing the strand discontinuities. On the other hand, E. coli MMR reads the absence of methylation as a strand discrimination signal. The MutS protein recognizes mismatches, In both MMR systems, strand discrimination is conducted by nicking endonucleases. MutL homologs from eukaryotes and most bacteria incise the discontinuous strand to introduce the entry or termination point for the excision reaction. In E. coli, MutH nicks the unmethylated strand of the duplex to generate the entry point of excision. Figure \(13\) shows different MMR pathway models. Figure \(13\): A schematic representation of MMR pathway models. Fukui, K. (2010) J. Nuc. Acids 260512. Creative Commons Attribution License Vertical panel (a): Eukaryotic MMR. A DNA mismatch is generated by the misincorporation of a base during DNA replication. MutSα recognizes base-base mismatches and MutLα nicks the 3-or5-side of the mismatched base on the discontinuous strand. The resulting DNA segment is excised by the EXO1 exonuclease, in cooperation with the single-stranded DNA-binding protein RPA. The DNA strand is resynthesized by DNA polymerase δ and DNA ligase 1. Vertical panel (b): MMR in mutH-less bacteria. Mismatched bases are recognized by MutS. After the incision of the discontinuous strand by MutL, the error-containing DNA strand is removed by the cooperative functions of DNA helicases, such as UvrD, the exonucleases RecJ and ExoI, and the single-stranded DNA-binding protein SSB. DNA polymerase III and DNA ligase fill the gap to complete the repair. Vertical panel (c): E. coli MMR. MutS recognizes mismatched bases, and MutL interacts with and stabilizes the complex. Then, MutH endonuclease is activated to incise the unmethylated GATC site to create an entry point for the excision reaction. DNA helicase, a single-stranded DNA-binding protein, and several exonucleases are involved in the excision reaction. PDB IDs of crystal structures in this figure are 2O8B (human MutSα), 1H7S (human MutLα), 1L1O (human RPA), 3IAY (human DNA polymerase δ), 1X9N (human DNA ligase 1), 1E3M (bacterial MutS), 1B63 (bacterial MutL), 2AZO (E. coli MutH), 2ISI (bacterial UvrD), 2ZXO (bacterial RecJ), 3C95 (bacterial ExoI), 2CWA (bacterial SSB), 2HQA (bacterial DNA polymerase III), and 2OWO (bacterial DNA ligase). Figure \(14\) shows an interactive iCn3D model of the E. Coli DNA Mismatch Repair Protein Muts Binding to a G-T Mismatch (1E3M). The MutS monomers are colored gray and cyan. The mismatched G9-T22 base pair is labeled. ADP is shown in spacefill. Phe 36 from the gray monomer is shown in magenta. The conformation of the monomers is different, so the dimer displays pseudo symmetry. Both subunits contribute to DNA binding, but only one (gray) binds both ADP and the actual mismatched GT base pair, the llatterthrough minor grove interactions, which kinks the DNA. General major grove interaction clamps the DNA. Note how far away the ADP binds. Phenylalanine 36 in the gray subunit (which binds the mismatch) inserts adjacent to the mismatch. ATP is bound and hydrolyzed to ADP by the MutS protein on binding the mismatch. Next, a dimer of MutL binds in a process that also requires ATP. MutH, a nuclease, also binds to MutL. The bound DNA is scanned until a "signal" is detected. In E. Coli, the signal is a GATC sequence that is methylated on just one strand and nicked by the MutH on the unmethylated GATC. Helicase II binds and unwinds the DNA in the region of the mismatch. Exonucleases (3' to 5' or 5' to 3') remove the sequence on the mismatched. PolII and DNA ligase then repair the DNA. MutS is yet another fascinating enzyme as it must scan millions of DNA bases without initiating repair until it localizes a mismatch. A series of sequential conformation changes that lead to specific recognition of the mismatch must occur. Fernandez-Leiro et al have determined the structure of MutS in a variety of stages along the repair pathway. Figure \(15\) shows interactive iCn3D models of the E. Coli MutS scanning form (EMD-11791, PDB 7AI5) and the more progressed MutS:MutL kink clamped form (EMD-11795, PDB 7AIC) E. Coli MutS scanning form (EMD-11791, PDB 7AI5) E. Coli MutS with MutL in kink clamped form (EMD-11795, PDB 7AIC) Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...pdHkXxzXsZnqu6 Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...hQ6PeKbQeCw916 Figure \(15\): E. Coli MutS scanning form (EMD-11791, PDB 7AI5) (left) and the more progressed MutS:MutL kink clamped form (EMD-11795, PDB 7AIC) (right) In the scanning form, the 2 monomers have been color coded as follows: monomer 1, N-terminal the interacts with DNA magenta, with the rest of the protein in red; monomer 2, the N-terminal part reacts with the DNA cyan, and the rest of the chain blue. In the kink-clamped state (right) the N-terminal magenta and N-terminal cyan sections were not present in the resolved structure. ATP is shown in spacefill. These structures suggest that during scanning by the homoduplex of normal DNA, the conformational change necessary for MutS to morph to the kink-clamped state can not occur due to a steric block. Kinking of the DNA at the mismatch removes the steric block. Click the links to download videos to get animations showing the role of MutS in mismatch repair. (Fernandez-Leiro, R., Bhairosing-Kok, D., Kunetsky, V. et al. The selection process of licensing a DNA mismatch for repair. Nat Struct Mol Biol 28, 373–381 (2021). https://doi.org/10.1038/s41594-021-00577-7, with permission) Video 1: Molecular mechanism of DNA mismatch repair initiation. Front and side views of MutS passing through the first four stages of the repair cascade: DNA scanning, mismatch recognition, intermediate state, and MutL recruitment. The movements show a computational morphing between the four cryo-EM structures. MutS monomer A is shown in a pale-green color, monomer B in pale blue, DNA in dark gray, and MutLLN40 in yellow. Video 2: Mismatch repair licensing at a mismatch. Top and side views of MutS as it transforms from the DNA-scanning state to the mismatch-bound state. The initial part of the movie represents the movement of monomer B relative to monomer A during the scanning state, as derived from the principal component analysis of the multibody refinement. Note that the MutS dimer explores multiple conformations, attempting to distort the DNA, without crossing over the opposite monomer. When a mismatch is present in the DNA, it allows MutS to deform and kink the DNA and the two MutS monomers to cross over in a clockwise manner. Movements show a computational morphing between the different states. MutS monomer A is shown in a pale-green color, monomer B in pale blue, and DNA in gray. The DNA mismatch is highlighted in pink Video 3: Multiple conformational changes of mismatch and connector domains tracking DNA. Front and side views of MutS as it goes from the mismatch-bound state to the MutLLN40-bound clamp state via the intermediate state. MutS monomer A is shown in a pale green color, monomer B in pale blue, and DNA in dark gray. DNA mismatch is highlighted in pink. The mismatch domain is shown in dark green and the connector domain in light green. The ends of a central helix in the connector domain are colored in red and blue for clarity. Movements show a computational morphing between the different states. Base Excision Repair Most oxidized bases are removed from DNA by enzymes operating within the Base Excision Repair (BER) pathway. Single-stranded DNA breaks can also be repaired through this process. Removal of oxidized bases in DNA is fairly rapid. For example, 8-oxo-dG was increased 10-fold in the livers of mice subjected to ionizing radiation, but the excess 8-oxo-dG was removed with a half-life of 11 minutes. 8-oxoG is excised by 8-oxoguanine DNA glycosylase (OGG1) leaving an apurinic site (AP site), as shown in Figure \(16\). AP sites are then processed further into single-strand breaks via backbone incision of AP-endonuclease 1 (APE1). In long patch base excision repair, the base and some additional nucleotides are replaced dependent on the activity of polymerase delta (Polδ) and epsilon (Polε) together with proliferating cell nuclear antigen (PCNA). The old strand is removed by Flap-endonuclease 1 (FEN1), before ligase I (LigI) ligates the backbone back together. Short patch base excision repair constitutes of polymerase beta (Polβ) replacing the single missing base, ligase III (LigIII) ligating the DNA backbone back together, and X-ray repair cross-complementing protein 1 (XRCC1) aiding the process and serving as a scaffold for additional factors. Base excision repair (BER) of 8-oxo-7,8-dihydroguanine (8-oxoG). Oxidative DNA damage is repaired via several repair intermediates by base excision repair (BER). Through the emoval of the oxidized base, a reactive apurinic site (AP site) is formed. Incision of the strand creates a single-strand break, and the damaged site is then repaired through either short or long patch BER. 25.2.6 Nucleotide Excision Repair Bulky DNA adducts and DNA crosslinks, such as those caused by UV light are repaired using Nucleotide Excision Repair (NER) pathways. In higher eukaryotic cells, NER excises 24-32 nucleotide DNA fragments containing the damaged lesion with extreme accuracy. Reparative synthesis using the undamaged strand as a template, followed by ligation of the single-strand break that emerged as a result of the damage, is the final stage of DNA repair. The process involves the coordinated action of approximately 30 proteins that successively form complexes with variable compositions on the DNA. NER consists of two pathways distinct in terms of initial damage recognition. Global genome nucleotide excision repair (GG-NER) detects and eliminates bulky damages in the entire genome, including the untranscribed regions and silent chromatin, while transcription-coupled nucleotide excision repair (TC -NER) operates when damage to a transcribed DNA strand limits transcription activity. TC-NER is activated by the stalling of RNA polymerase II at the damaged sites of a transcribed strand, while GG-NER is controlled by the protein, XPC, a specialized protein factor that reveals the damage. A schematic GG-NER process is presented in Figure (17\) below. Genetic mutations in NER pathway genes can result in UV-sensitive and high-carcinogenic pathologies, such as xeroderma pigmentosum (XP), Cockayne syndrome (CS), and trichothiodystrophy (TTD), as well as some neurodegenerative manifestations. Xeroderma pigmentosum has provided the names of some of the genes involved in NER. Mutation of XP genes and loss of proper NER function causes the symptoms associated with the disease. People with XP have an impaired ability to repair bulky DNA adducts and crosslinks, such as thymine dimers that are caused by UV-light exposure. People suffering from XP have extreme photosensitivity, skin atrophy, hyperpigmentation, and a high rate of sunlight-induced skin cancer. The risk of internal tumors in XP patients is also 1,000-fold higher. Moreover, the disease is often associated with neurologic disorders. Currently, there is no effective treatment for this disorder. The detection of bulky DNA lesions during NER is particularly challenging for a cell, which can be solved only through highly sensitive recognition that requires multiple protein components. In contrast to BER, where a damaged base is simultaneously recognized and eliminated by a single specialized glycosylase, specialized groups of proteins are responsible for the recognition of the lesion and the excision of the lesion in NER. In eukaryotic NER, universal sensor proteins perform the initial recognition of the total range of bulky damages. In the case of TC-NER, it occurs when the transcribing RNA polymerase II is stalled by damage; in GG-NER, these are complexes of the XPC factor and DDB1-DDB2 heterodimer (XPE factor) enhancing the repair of UV damage. In general, NER recognition of damage is a multistep process involving several proteins that form near damaged complexes of variable compositions. The process is completed by the formation of a preincision complex ready to eliminate a damaged DNA fragment by specialized NER endonucleases. In a eukaryotic cell after stable XPC/DNA complex formation during the initial recognition of the damage, NER is performed by a repairasome, which is a complex of variable composition and architecture consisting of a large number of subunits. Individual subunits of the complex have no sufficient affinity and selectivity to the substrate (DNA containing bulky damage). The situation changes when specific protein complexes are established at the damage site. A total of 18 polypeptides must be accurately positioned within two or three DNA turns when a stable structure ready for damage removal is formed and excision starts. The structure of NER-associated proteins provides the possibility of contact with the DNA substrate and of dynamic specific protein-protein interactions. The changes in interactions performed by the same protein are one of the mechanisms that regulate the repair process and fine-tune the complexes, providing high-precision nucleotide excision repair. 25.2.7 Repair of Double-Stranded DNA Breaks Cells have evolved two main pathways to repair double-strand breaks within the DNA: the non-homologous end-joining (NHEJ) pathway, which ensures direct resealing of DNA ends; and the homologous recombination (HR) pathway that relies on the presence of homologous DNA sequences for DSB repair, as shown in Figure (18\) below. NHEJ repair is the simplest and most widely utilized mechanism to repair DSB that occur in DNA. Repair by NHEJ involves direct resealing of the two broken ends independently of sequence homology. Although being active throughout the cell cycle, NHEJ is relatively more important during the G1 phase. Proteins required for NHEJ include but are not restricted to, the highly conserved Ku70/Ku80 heterodimeric complex, DNA-dependent protein kinase catalytic subunit (DNA-PKcs), and DNA Ligase IV (LIG4) in complex with XRCC4. By directly binding DNA ends, Ku70/Ku80 ensures protection against exonucleases and, as such, acts as an inhibitor of HR. Very short sequence homologies are likely to help DNA end alignment before NHEJ-dependent repair, however, they are not strictly required. NHEJ protects genetic integrity by rejoining broken strands of DNA that may otherwise be lost during DNA replication and cell regeneration. However, during the process of NHEJ, insertions or deletions within the joined regions may occur (Fig 25.2.17). Non-homologous end-joining (NHEJ) and homologous recombination (HR) pathways act competitively to repair DNA double-strand breaks (DSBs). Key players of NHEJ and HR are depicted. The MRE11/RAD50/XRS2 (MRX) complex is recruited very early at DNA ends and appears to play important roles for both NHEJ and HR. Ku70/Ku80 heterodimer is required for NHEJ and, through inhibition of DNA end resection (5′–3′ exo), acts as a repressor of HR. The fidelity of NHEJ-dependent DSB repair is low and, most of the time, associated with nucleotide deletions and/or insertions at repair junctions. The common early step of HR-dependent mechanisms is the formation of ssDNA which is then coated by replication protein A (RPA). Single-strand annealing (SSA) mechanism requires the presence of direct repeats (shown in orange) on both sides of the break. SSA does not imply any strand invasion process and is therefore not dependent on RAD51 protein. Strand invasion and D-loop formation are however common steps of synthesis-dependent strand annealing (SDSA) and double Holliday junction (HJ) dissolution mechanisms. In the latter case, double Holliday junctions are resolved with or without crossing over. In contrast to NHEJ, homologous recombination (HR) requires a homologous DNA sequence to serve as a template for DNA-synthesis-dependent repair and involves extensive DNA-end processing. As expected, HR is extremely accurate, as it leads to precise repair of the damaged locus using DNA sequences homologous to the broken ends. HR predominantly uses the sister chromatid as a template for DSB repair, rather than the homologous chromosome. Correspondingly, HR is largely inhibited while cells are in the G1 phase of the cell cycle when the sister chromatid has not yet been replicated, as shown in panel (A) of Figure (19\) below. HR repair mechanisms play a bigger role in DSB repair that occurs after S-phase DNA replication (S-phase, G2, and M). Repair through HR is not defined by a unique mechanism but operates through various mechanistically distinct DSB repair processes, including synthesis-dependent strand annealing (SDSA), double Holliday junction resolution, and single-strand annealing (SSA). The common step for HR-dependent DSB repair mechanisms is the initial formation of single-stranded DNA (ssDNA) for pairing with homologous DNA template sequences. For this to occur, the 5' DNA strand at the DSB is processed by multiple nucleases and accessory proteins to create a 3' ssDNA section that can be used as a template for recombination (see Figure 18 above). Panel B of Figure (19\) below provides a more detailed look at the HR process. During the highly regulated process of HR, three main phases can be distinguished. Firstly, 3′-single-stranded DNA (ssDNA) ends are generated by nucleolytic degradation of the 5′-strands. This first step is catalyzed by endonucleases, including the MRN complex (consisting of Mre11, Rad50, and Nbs1). In the second step, the ssDNA-ends are coated by replication protein A (RPA) filaments. In the third step, RPA is replaced by Rad51 in a BRCA1- and BRCA2-dependent process, to ultimately perform the recombinase reaction using a homologous DNA template. Importantly, HR is not only employed to repair DNA lesions induced by DNA-damaging agents but is also essential for proper chromosome segregation during meiosis. The relevance of HR in these physiological processes is illustrated by its strict requirement during development. Mice lacking key HR genes, such as Brca1, Brca2, or Rad51, display extensive genetic alterations which lead to early embryonic lethality. Whereas homozygous inactivation of HR genes is usually embryonic lethal, heterozygous inactivation of,BRCA1 and BRCA2, does not interfere with cellular viability but rather predisposes individuals to cancer, including breast and ovarian cancer. The tumors that develop in individuals with heterozygous BRCA1/2 mutations invariably lose their second BRCA1/2 allele, indicating that in certain cancers, the absence of BRCA1/2 is compatible with cellular proliferation. How exactly such tumors cope with their HR defect is currently not fully understood. Panel (A) shows the DNA DSBs repair pathways in the context of cell cycle regulation. Non-homologous end joining (NHEJ) can be performed throughout the cell cycle and is indicated with the red line. Homologous recombination (HR) can only be employed in S/G2 phases of the cell cycle and is indicated in green. Pane (B) shows the key steps in the HR repair pathway are indicated. After DSB recognition, 5′–3′ end resection is initiated by the MRN (Mre11, Rad50, Nbs1) complex and CtIP. Subsequently, further resection by the Exo1, DNA2, and Sgs1 proteins is conducted to ensure ‘maintained’ resection. Then, resected DNA ends are bound by replication protein A (RPA). The actual recombination step within HR repair, termed strand exchange, is executed by the recombinase Rad51. Rad51 replaces RPA to eventually assemble helical nucleoprotein filaments called ‘presynaptic filaments.’ This process is facilitated by other HR components, including BRCA1 and BRCA2. The final step of junction resolution is executed by helicases including Bloom syndrome, the RecQ helicase-like (BLM) helicase. Error-Prone Bypass and Translesion Synthesis If DNA is not repaired before DNA replication, the cell must employ another strategy to replicate the DNA, even in the presence of a DNA lesion. This is important to avoid causing double-stranded DNA breaks that can occur when a replisome stalls at the replication fork. Under these circumstances, another strategy that cells use to respond to DNA damage is to bypass lesions found during DNA replication and continue with the replicative process. DNA damage bypass can occur by recombination mechanisms or through a novel mechanism called translesion synthesis. Translesion synthesis employs an alternate DNA polymerase that can substitute for a DNA polymerase that has stalled at the replication fork due to DNA damage. Specialized DNA polymerases, that are active in regions with DNA damage, have active sites that can accommodate fluctuations in DNA topography that enable them to bypass the lesions and continue with the replicative process. The evolution of DNA polymerases that can tolerate the presence of distorted DNA lesions and continue with the replicative process can be seen at all levels of life, from prokaryotic, single-celled organisms through eukaryotic multicellular organisms, including humans. In fact, within vertebrates, there has been a large expansion of DNA polymerases that play a role in DNA damage bypass mechanisms and highlight the importance of these processes in damage tolerance and cell survival, as shown in Table (3\) below. Table (3\): DNA Polymerases involved in Error-Prone Bypass The activity of error-prone DNA polymerases is tightly regulated to avoid the rampant introduction of mutations within the DNA sequence. One of the main mechanisms that is employed within a replisome that is stalled at the replication fork due to DNA damage, involves the monoubiquitination of PCNA. Recall from Chapter 9, that PCNA is the sliding clamp that enables the DNA polymerase to bind tightly enough with the DNA during replication to mediate efficient DNA synthesis. Monoubiquitination of PCNA enables the recruitment of a translesion DNA polymerases and the bypass of the damaged lesions during DNA synthesis. During translesion synthesis, the polymerase must insert a dNTP opposite of the lesion. None of the dNTP bases will likely be able to form stable hydrogen bond interactions with the damaged lesion. Thus, the nucleotide that causes the least distortion or repulsion will usually be added across from the lesion. This can cause transition or transversion mutations to occur at the lesion location. Alternatively, translesion polymerases can be prone to slippage, and either causes an insertion or deletion mutation in the vicinity of the DNA lesion. These slippages can lead to frameshift mutations if they occur within gene coding regions. Thus, over a lifetime, translesion synthesis in multicellular organisms can lead to an accumulation of mutations within somatic cells and cause the formation of tumors and the disease of cancer. Evolution by natural selection is also possible due to random mutations that occur within germ cells. Occasionally, germline mutations may lead to a beneficial mutation that enhances the survival of an individual within a population. If this gene proves to enhance the survival of the population, it will be selected over time within the population and cause the evolution of that species. An example of a beneficial mutation is the case of a population of people that show resistance to HIV infection. Since the first case of infection with human immunodeficiency virus (HIV) was reported in 1981, nearly 40 million people have died from HIV infection, the virus that causes acquired immune deficiency syndrome (AIDS). The virus targets helper T cells that play a key role in bridging the innate and adaptive immune response, infecting and killing cells normally involved in the body’s response to infection. There is no cure for HIV infection, but many drugs have been developed to slow or block the progression of the virus. Although individuals around the world may be infected, the highest prevalence among people 15–49 years old is in sub-Saharan Africa, where nearly one person in 20 is infected, accounting for greater than 70% of the infections worldwide, as shown in Figure (20\) below. Unfortunately, this is also a part of the world where prevention strategies and drugs to treat the infection are the most lacking. In recent years, scientific interest has been piqued by the discovery of a few individuals from northern Europe who are resistant to HIV infection. In 1998, American geneticist Stephen J. O’Brien at the National Institutes of Health (NIH) and colleagues published the results of their genetic analysis of more than 4,000 individuals. These indicated that many individuals of Eurasian descent (up to 14% in some ethnic groups) have a deletion mutation, called CCR5-delta 32, in the gene encoding CCR5. CCR5 is a coreceptor found on the surface of T-cells that is necessary for many strains of the virus to enter the host cell. The mutation leads to the production of a receptor to which HIV cannot effectively bind and thus blocks viral entry. People homozygous for this mutation have greatly reduced susceptibility to HIV infection, and those who are heterozygous have some protection from infection as well. It is not clear why people of northern European descent, specifically, carry this mutation, but its prevalence seems to be highest in northern Europe and steadily decreases in populations as one moves south. Research indicates that the mutation has been present since before HIV appeared and may have been selected for in European populations as a result of exposure to the plague or smallpox. This mutation may protect individuals from the plague (caused by the bacterium Yersinia pestis) and smallpox (caused by the variola virus) because this receptor may also be involved in these diseases. The age of this mutation is a matter of debate, but estimates suggest it appeared between 1875 years to 225 years ago, and may have been spread from Northern Europe through Viking invasions. This exciting finding has led to new avenues in HIV research, including looking for drugs to block CCR5 binding to HIV in individuals who lack the mutation. Although DNA testing to determine which individuals carry the CCR5-delta 32 mutation is possible, there are documented cases of individuals homozygous for the mutation contracting HIV. For this reason, DNA testing for the mutation is not widely recommended by public health officials so as not to encourage risky behavior in those who carry the mutation. Nevertheless, inhibiting the binding of HIV to CCR5 continues to be a valid strategy for the development of drug therapies for those infected with HIV. Practice Problems Multiple Choice Which of the following is a change in the sequence that leads to the formation of a stop codon? 1. missense mutation 2. nonsense mutation 3. silent mutation 4. deletion mutation [reveal-answer q=”745512″]Show Answer[/reveal-answer] [hidden-answer a=”745512″]Answer b. A nonsense mutation is a change in the sequence that leads to formation of a stop codon.[/hidden-answer] The formation of pyrimidine dimers results from which of the following? 1. spontaneous errors by DNA polymerase 2. exposure to gamma radiation 3. exposure to ultraviolet radiation 4. exposure to intercalating agents [reveal-answer q=”709151″]Show Answer[/reveal-answer] [hidden-answer a=”709151″]Answer c. The formation of pyrimidine dimers results from exposure to ultraviolet radiation.[/hidden-answer] Which of the following is an example of a frameshift mutation? 1. a deletion of a codon 2. missense mutation 3. silent mutation 4. deletion of one nucleotide [reveal-answer q=”688366″]Show Answer[/reveal-answer] [hidden-answer a=”688366″]Answer a. The deletion of one nucleotide is an example of a frameshift mutation.[/hidden-answer] Which of the following is the type of DNA repair in which thymine dimers are directly broken down by the enzyme photolyase? 1. direct repair 2. nucleotide excision repair 3. mismatch repair 4. proofreading [reveal-answer q=”755583″]Show Answer[/reveal-answer] [hidden-answer a=”755583″]Answer a. In a direct repair, thymine dimers are directly broken down by the enzyme photolyase.[/hidden-answer] Which of the following regarding the Ames test is true? 1. It is used to identify newly formed auxotrophic mutants. 2. It is used to identify mutants with restored biosynthetic activity. 3. It is used to identify spontaneous mutants. 4. It is used to identify mutants lacking photoreactivation activity. [reveal-answer q=”770537″]Show Answer[/reveal-answer] [hidden-answer a=”770537″]Answer b. It is used to identify mutants with restored biosynthetic activity.[/hidden-answer] Fill in the Blank A chemical mutagen that is structurally similar to a nucleotide but has different base-pairing rules is called a ________. [reveal-answer q=”702924″]Show Answer[/reveal-answer] [hidden-answer a=”702924″]A chemical mutagen that is structurally similar to a nucleotide but has different base-pairing rules is called a nucleoside analog.[/hidden-answer] The enzyme used in light repair to split thymine dimers is called ________. [reveal-answer q=”939657″]Show Answer[/reveal-answer] [hidden-answer a=”939657″]The enzyme used in light repair to split thymine dimers is called photolyase.[/hidden-answer] The phenotype of an organism that is most commonly observed in nature is called the ________. [reveal-answer q=”640686″]Show Answer[/reveal-answer] [hidden-answer a=”640686″]The phenotype of an organism that is most commonly observed in nature is called the wild type.[/hidden-answer] True/False Carcinogens are typically mutagenic. [reveal-answer q=”166576″]Show Answer[/reveal-answer] [hidden-answer a=”166576″]True[/hidden-answer] Think about It Why is it more likely that insertions or deletions will be more detrimental to a cell than point mutations? Critical Thinking Below are several DNA sequences that are mutated compared with the wild-type sequence: 3′-T A C T G A C T G A C G A T C-5′. Envision that each is a section of a DNA molecule that has separated in preparation for transcription, so you are only seeing the template strand. Construct the complementary DNA sequences (indicating 5′ and 3′ ends) for each mutated DNA sequence, then transcribe (indicating 5′ and 3′ ends) the template strands, and translate the mRNA molecules using the genetic code, recording the resulting amino acid sequence (indicating the N and C termini). What type of mutation is each? Mutated DNA Template Strand #1: 3′-T A C T G T C T G A C G A T C-5′ Complementary DNA sequence: [practice-area rows=”1″][/practice-area] mRNA sequence transcribed from template: [practice-area rows=”1″][/practice-area] Amino acid sequence of peptide: [practice-area rows=”1″][/practice-area] Type of mutation: [practice-area rows=”1″][/practice-area] Mutated DNA Template Strand #2: 3′-T A C G G A C T G A C G A T C-5′ Complementary DNA sequence: [practice-area rows=”1″][/practice-area] mRNA sequence transcribed from template: [practice-area rows=”1″][/practice-area] Amino acid sequence of peptide: [practice-area rows=”1″][/practice-area] Type of mutation: [practice-area rows=”1″][/practice-area] Mutated DNA Template Strand #3: 3′-T A C T G A C T G A C T A T C-5′ Complementary DNA sequence: [practice-area rows=”1″][/practice-area] mRNA sequence transcribed from template: [practice-area rows=”1″][/practice-area] Amino acid sequence of peptide: [practice-area rows=”1″][/practice-area] Type of mutation: [practice-area rows=”1″][/practice-area] Mutated DNA Template Strand #4: 3′-T A C G A C T G A C T A T C-5′ Complementary DNA sequence: [practice-area rows=”1″][/practice-area] mRNA sequence transcribed from template: [practice-area rows=”1″][/practice-area] Amino acid sequence of peptide: [practice-area rows=”1″][/practice-area] Type of mutation: [practice-area rows=”1″][/practice-area] <h212references">25.2.10 References 1. <lifootnote-99-2">World Health Organization. " Global Health Observatory (GHO) Data, HIV/AIDS." http://www.who.int/gho/hiv/en/. Accessed August 5, 2016. 2. Parker, N., Schneegurt, M., Thi Tu, A-H., Lister, P., Forster, B.M. (2019) Microbiology. Openstax. Available at: https://opentextbc.ca/microbiologyopenstax/ 3. Wikipedia contributors. (2020, July 15). DNA oxidation. In Wikipedia, The Free Encyclopedia. Retrieved 03:44, July 16, 2020, from https://en.Wikipedia.org/w/index.php?title=DNA_oxidation&oldid=967811859 4. Wakim, S. and Grewal, M. (2020) Human Biology. Libretexts. Available at: https://bio.libretexts.org/Bookshelves/Human_Biology/Book%3A_Human_Biology_(Wakim_and_Grewal) 5. Ahmad, A., Nay, S.L. and O'Conner, T.R. (2015) Chapter 4 Direct Reversal Repair in Mammalian Cells. Published through INTECH. Available at: https://cdn.intechopen.com/pdfs/48191.pdf 6. Wikipedia contributors. (2020, April 20). AP site. In Wikipedia, The Free Encyclopedia. Retrieved 18:15, July 23, 2020, from https://en.Wikipedia.org/w/index.php?title=AP_site&oldid=952117602 7. Wikipedia contributors. (2020, July 4). Benzo(a)pyrene. In Wikipedia, The Free Encyclopedia. Retrieved 06:15, July 24, 2020, from https://en.Wikipedia.org/w/index.php?title=Benzo(a)pyrene&oldid=965990545 8. Wikipedia contributors. (2020, June 23). Pyrimidine dimer. In Wikipedia, The Free Encyclopedia. Retrieved 06:54, July 24, 2020, from https://en.Wikipedia.org/w/index.php?title=Pyrimidine_dimer&oldid=964108515 9. Morimoto, S., Tsuda, M., Bunch, H., Sasanuma, H., Ausin, C. and Takeda, S. (2019) Type II DNA Topoisomerases Cause Spontaneous Double-Strand Breaks in Genomic DNA. Genes 10:868. Available at: https://www.researchgate.net/publication/336916880_Type_II_DNA_topoisomerases_cause_spontaneous_double-strand_breaks_in_genomic_DNA/figures?lo=1 10. Ding, L., Cao, J., Lin, W., Chen, H., Xiong, X., Ao, H., Yu, M., Lin, J., Cui, Q. (2020) The Roles of the Cyclin-Dependent Kinases in Cell-Cycle Progression and Therapeutic Strategies in Human Breast Cancer. Int. J. Mol Sci 21(6):1960. Available at: https://www.mdpi.com/1422-0067/21/6/1960/htm 11. Verma, N., Franchitto, M., Zonfrilli, A., Cialfi, S., Palermo, R., and Talora, C. (2019) DNA Damage Stress: Cui Prodest? Int. J. Mol. Sci. 20(5):1073. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC6429504/ 12. Fukui, K. (2010) DNA Mismatch Repair in Eukaryotes and Bacteria. J. Nuc. Acids 260512. Available at: https://www.hindawi.com/journals/jna/2010/260512/#copyright 13. Petruseva, I.O., Evdokimov, A.N., and Lavrik, O.I. (2014) Molecular Mechanism of Global Genome Nucleotide Excision Repair. Acta Naturae 6(1):23-34. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC3999463/ 14. Decottingnies, A. (2013) Alternative end-joining mechanisms: A historical perspective. Frontiers in Genetics 4(48):48. Available at: https://www.researchgate.net/publication/236129718_Alternative_end-joining_mechanisms_A_historical_perspective 15. Vitor, A.C., Huertas, P., Legube, G., and de Almeida, S.F. (2020) Studying DNA Double-Strand Break Repair: An Every-Growing Toolbox. Front. Mol Biosci 7:24. Available at: https://www.frontiersin.org/articles/10.3389/fmolb.2020.00024/full 16. Krajewska, M., Fehrmann, R.S.N., de Vries, E.G.E., and van Vugt, A.A.T.M. (2015) Regulators of homologous recombination repair as novel targets for cancer treatment. Front. Genet. 6:96. Available at: https://www.frontiersin.org/articles/10.3389/fgene.2015.00096/full

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/24%3A_DNA_Metabolism/24.02%3A_DNA_Mutations_Damage_and_Repair.txt

Search Fundamentals of Biochemistry Homologous Recombination Homologous recombination is a type of genetic recombination in which genetic information is exchanged between two similar or identical molecules of double-stranded or single-stranded nucleic acids (usually DNA as in cellular organisms but may be also RNA in viruses). As noted in section 25.2, This process is widely used by cells to accurately repair harmful breaks that occur on both strands of DNA, known as double-strand breaks (DSB), in a process called homologous recombinational repair (HRR). Homologous recombination also produces new combinations of DNA sequences during meiosis, the process by which eukaryotes make gamete cells, like sperm and egg cells in animals. These new combinations of DNA represent genetic variation in offspring, which in turn enables populations to adapt during evolution. Homologous recombination is also used in horizontal gene transfer to exchange genetic material between different strains and species of bacteria and viruses. Although homologous recombination varies widely among different organisms and cell types, for double-stranded DNA (dsDNA) most forms involve the same basic steps. After a double-strand break occurs, sections of DNA around the 5' ends of the break are cut away in a process called resection. In the strand invasion step that follows, an overhanging 3' end of the broken DNA molecule then "invades" a similar or identical DNA molecule that is not broken. After strand invasion, the further sequence of events may follow either of two main pathways discussed below (see Models); the DSBR (double-strand break repair) pathway or the SDSA (synthesis-dependent strand annealing) pathway. Homologous recombination that occurs during DNA repair tends to result in non-crossover products, in effect restoring the damaged DNA molecule as it existed before the double-strand break. There are several different ways to repair DSB as illustrated in Figure \(1\). The broken (or resected) DNA with a double-stranded break must find and come together (synapse), invade (or intertwine) with the DNA of a donor, that is homologous to it. The repair can then ensue. In somatic cells that undergo mitosis and not meiosis, the preferred donor is the sister chromatid (the copy of one chromosome made during cell division) and not the homologous chromosome (from the diploid cell). Variants include synthesis-dependent strand annealing pathway (SDSA). Other variants include nonhomologous end-joining (NHEJ), microhomology-mediated end-joining (MMEJ), and double Holiday junction (dHJ). Figure \(1\): Model for the repair of DNA double-strand breaks by homologous recombination in somatic cells. Wright et al. J. Biol. Chem. (2018) 293(27) 10524 –10535. Creative Commons Attribution (CC BY 4.0) When a DNA double-strand break (DSB) occurs in a DNA molecule, a repair can proceed by multiple pathways largely controlled by end resection. NHEJ is capable of repairing unresected or minimally resected DSBs in a template-independent fashion. MMEJ and single-strand annealing (SSA) rely on different extents of homology between the two DSB ends for repair independent of a donor molecule. Homologous recombination proceeds as shown in the figure using a homologous donor DNA. Most of the extended D-loops in somatic cells are disrupted and subsequently repaired by SDSA. The result of the repair by SDSA is always a noncrossover outcome, thus avoiding the loss of heterozygosity produced by somatic crossovers. SDSA occurs by disruption of the extended D-loop and annealing the newly synthesized DNA with the second end of the broken molecule. Alternatively, the newly synthesized strand may invade the second end. The extended D-loop can also undergo second-end capture or invasion to form a double Holliday junction (dHJ). This may either lead to a crossover or a noncrossover outcome. Invasion by the second break end makes dHJ formation and hence crossover outcome more likely for another model for crossover generation. dHJs can be dissolved into noncrossovers by the concerted action of the Sgs1–Top3–Rmi1 complex to migrate the two junctions toward each other and then decatenate the strands of the hemicatenane by the Top3 topoisomerase activity. Each colored line indicates a strand of DNA, and dotted lines represent DNA synthesis. In the process of homologous recombination, a key intermediate is the Holiday junction, named after Robin Holiday who discovered it. It consists of branched nucleic acid with four double-stranded arms joined. A holiday junction is seen as the crossing of red and blue strands in the middle of Figure 1 labeled Nascent D-loop. Also, one is seen in the Extended D-loop just below it. A double Holiday junction is seen in the middle of the right-hand section. Two views of Holiday junctions are shown in Figure \(2\). By Донор - Own work, CC BY-SA 4.0, https://commons.wikimedia.org/w/inde...curid=48470765 By Antony-22 - Own work, CC BY-SA 4.0, https://commons.wikimedia.org/w/inde...curid=38557614 Figure \(2\). Two views of holiday junctions. The left-hand panel shows the primary and secondary sequences and some tertiary (3D) aspects of base-stacking conformational isomers of the Holliday junction. The bases nearest to the junction point determine which stacked isomer dominates. Figure \(3\) shows an interactive iCn3D model of the structure of the Holliday junction intermediate in Cre-loxP site-specific recombination (3CRX). The alpha carbon backbone of the four Cre recombinase monomers in the tetramer is shown in red. The DNA is nearly planar with a twofold-symmetric DNA intermediate that is similar to a square and stacked Holiday junction for the DNA in the unbound state. Homologous recombination is conserved across all three domains of life as well as DNA and RNA viruses, suggesting that it is a nearly universal biological mechanism. The discovery of genes for homologous recombination in protists—a diverse group of eukaryotic microorganisms—has been interpreted as evidence that meiosis emerged early in the evolution of eukaryotes. Since their dysfunction has been strongly associated with increased susceptibility to several types of cancer, the proteins that facilitate homologous recombination are topics of active research. Homologous recombination is also used in gene targeting, a technique for introducing genetic changes into target organisms. For their development of this technique, Mario Capecchi, Martin Evans, and Oliver Smithies were awarded the 2007 Nobel Prize for Physiology or Medicine; Capecchi[3] and Smithies[4] independently discovered applications to mouse embryonic stem cells, however, the highly conserved mechanisms underlying the DSB repair model, including uniform homologous integration of transformed DNA (gene therapy), were first shown in plasmid experiments by Orr-Weaver, Szostack, and Rothstein. Before the beginning of meiosis, the replication of the DNA is required to form sister chromatids, as shown in Step 1 in Figure \(4\). Once replicated, the DNA will condense and begin the process of meiosis. As the cells enter metaphase of meiosis, heterologous chromosomes are paired together (Step 2). When the homologous chromosomes are paired, they can under genetic mixing or DNA recombination form new chromosomal arrangements that are unique from the parental chromosomes (Step 3). Once the process of recombination is finished, the heterologous chromosomes are separated into two different daughter cells (Step 4). This is called Meiosis I. At this stage the chromosomes have been reduced from the diploid to the haploid state, however, each chromosome set is still paired with its sister chromatid and needs to undergo a second round of cell division to produce the final set of four gametes (Step 5). This is called Meiosis II and results in the formation of four haploid gametes that are genetically unique. During the process of meiosis, cell division is used to create the gametes or reproductive cells of an organism (the egg and the sperm cells). Meiosis results in the reduction of the genome from the 2n or diploid state to the 1n or haploid state. As you can see in Figure 25.3.1, the process of meiotic division results in the generation of four genetically unique haploid cells and involves the pairing of heterologous chromosomes during metaphase of meiosis. In humans, the meiotic process results in four viable sperm cells in the male and a single viable egg in the female. The other three cells produced in the female during the meiotic division are termed polar bodies and are very small and do not contain enough cytoplasmic components to survive. They get reabsorbed into the body. In either case, the resulting egg or sperm cell each carries a single copy of the genome and is in the haploid state. It is during the process of meiosis that homologous recombination occurs in a controlled manner to introduce genetic variation into the resulting gametes. As a result, each egg and sperm cell has a unique genetic makeup that is a mixture of both parental copies of the genome. Proper segregation during meiosis requires that homologs be connected by the combination of crossovers and sister chromatid cohesion. To generate crossovers, numerous double-strand breaks (DSBs) are introduced throughout the genome by the conserved Spo11 endonuclease. DSB formation and its repair are then highly regulated to ensure that homologous chromosomes contain at least one crossover and that no DSBs remain before meiosis I segregation. The synaptonemal complex (SC) is a meiosis-specific structure formed between homologous chromosomes during prophase that promotes DSB formation and biases the repair of DSBs to homologous chromosomes rather than back to the sister chromatids, ensuring that genetic recombination occurs. Synapsis, the pairing of homologous chromosomes, occurs when a particular recombination pathway is successful in establishing stable interhomolog connections. Formation of the Synaptonemal Complex In the 1950s, electron microscopists discovered an evolutionarily conserved, meiosis-specific structure formed between homologous chromosomes, unique to prophase I, called the SC, as shown in Figure \(5\). The SC physically connects homologs during prophase I and is removed before metaphase I, when homologs are connected instead by the combination of crossovers and sister chromatid cohesion. What is the function of the SC? Decades of research have shown that this elaborate chromosomal structure is critical for the regulation of recombination, the process by which crossovers are generated. Panel (A) shows the synaptonemal complex. When chromosomes form synapses, recombination intermediates contain double Holliday junctions (shown by intersecting loops). When cells exit the pachytene stage, the stage of meiotic prophase when chromosomes fully synapse, Holliday junctions are resolved to form crossovers, and the SC is disassembled. DSB formation is greatly reduced by synapsis but is not completely abolished until cells exit pachynema. Panel (B) shows the sequence diversity between homologous chromosomes largely inhibits recombination and synapsis, resulting in persistent DSB formation. Panel (C) shows that in the absence of the central element, the transverse filament is not assembled, resulting in chromosomes that lack the central region. Recombination intermediates containing double Holliday junctions are still formed, but DSB formation is not down-regulated. Image from: SC formation begins with the condensation of sister chromatids along meiosis-specific protein cores to make axial elements. Axial elements from homologous chromosomes are “zippered” together by the insertion of the central region. (Note that after synapsis, axial elements are called lateral elements (Panel A above). The central region is comprised of (1) transverse filaments located perpendicular to the lateral elements, and (2) the central element, which runs parallel to the lateral elements midway through the central region. Assembly of the SC is initiated in the early stage of meiotic prophase I, which is commonly divided into five substages (leptotene, zygotene, pachytene, diplotene, and diakinesis). For proper assembly of the SC followed by the correct pairing of the homologous chromosome, lateral elements (LEs), which are composed of two main proteins (SYCP2 and SYCP3) should be formed along each chromosome at the initial stage, during leptotene. Later the two LEs associate with the linker part, known as the transverse filaments (TFs). TFs are primarily composed of the protein SYCP1. The central element (CE), which is composed of SYCE1, SYCE2, SYCE3, and TEX12, then connects to the LEs through the TFs, as shown in Figure \(6\). The lateral elements complete their pairing during the zygotene stage leading to the formation of the tripartite SC structure seen during the pachytene stage of the first meiotic prophase as shown in Figure \(7\) and Figure \(8\). This occurs in both males and females during gametogenesis. Panel (A) shows a model of the SC. Lateral elements (light blue rods) of homologous chromosomes align and synapse together via a meshwork of transverse filaments (black lines) and longitudinal filaments (dark blue rods). The longitudinal filaments are collectively referred to as the “central element” of the SC. Ellipsoidal structures called recombination nodules (gray ellipsoid) are constructed on the central region of the SC. As their name implies, recombination nodules are believed to be involved in facilitating meiotic recombination (crossing over). The chromatin (red loops) of each homolog is attached to its corresponding lateral element. Because there are two “sister chromatids” in each homolog, two loops are shown extending laterally from each point along a lateral element. Panel (B) Top shows a set of tomato SCs. Chromatin “sheaths” are visible around each SC showing two tomato SCs. The chromatin has been stripped from the SCs, allowing the details of the SC to be observed. Each SC has a kinetochore (“ball-like” structure) at its centromere. Recombination nodules, ellipsoidal structures found on the central regions of SCs, mark the sites of crossover events (see inset). Zygotene is the sub-stage where synapsis between homologous chromosomes begins. It is also known as zygonema. This synapse can form up and down the chromosomes allowing numerous points of contact called 'synaptonemal complex', this can be compared to a zipper structure, due to the coils of chromatin. The SC facilitates synapsis by holding the aligned chromosomes together. After the homologous pairs synapse they are either called tetrads or bivalents. Bivalent is more commonly used at an advanced level as it is a better choice due to similar names for similar states (a single homolog is a 'univalent', and three homologs are a 'trivalent'). Once the synapse is formed it is called a bivalent (where a chromatid of one pair is synapsed/attached to the chromatid in a homologous chromosome and crossing over can occur. Subsequently, the synapses snap completing the crossing over of the genetic information. As a result, the variation in genetic material has increased significantly, because up and down the chromosome there has been an exchange of the mother and father's genetic material. The two sister chromatids separate from each other, but the homologous chromosomes remain attached. This makes the complex look much thicker. The SC is complete, allowing chiasma to form. This is what allows the crossing over alleles to occur as this is a process that only happens over a small region of the chromosomes. The chiasma is a structure that forms between a pair of homologous chromosomes by crossover recombination and physically links the homologous chromosomes during meiosis as shown in Figure \(9\). Chiasmata are essential for the attachment of the homologous chromosomes to opposite spindle poles (bipolar attachment) and their subsequent segregation to the opposite poles during meiosis I. Mechanism of Homologous Recombination Meiotic recombination is a tightly regulated process that is triggered by the programmed induction of DNA double-strand breaks (DSBs). Once formed, the ends of the DSBs are nucleolytically processed to generate 3′ single-stranded DNA (ssDNA) tails. Meiotic recombination factors then engage these ssDNA tails to form a nucleoprotein ensemble capable of locating DNA homology in the chromosome homolog and mediating invasion of the homolog to form a DNA joint called a displacement loop or D-loop. The 3′ end of the invading strand is extended by DNA synthesis, followed by the pairing of the non-invading 3′ single-stranded tail with the displaced ssDNA strand in the enlarged D-loop (second-end capture). After DNA synthesis and DNA ligation, a double Holliday Junction (dHJ) intermediate is formed. Resolution of the dHJ intermediate can result in crossover recombinants that harbor a reciprocal exchange of the arms of the homologous chromosomes. Genetic studies have revealed that meiotic DSBs arise via the action of a protein ensemble that harbors the Spo11 protein, which bears homologous to archaeal Topo VIA, the catalytic subunit of a type II topoisomerase. Indeed, studies in S. cerevisiae, S. pombe, and M. musculus have shown that Spo11 becomes covalently conjugated to the 5′ ends of DNA through a tyrosine residue proposed to be the catalytic center of topoisomerase function. Thus, mutations in the putative catalytic tyrosine residue of Spo11 engender the same phenotype as spo11 deletion in S. cerevisiae, S. pombe, A. thaliana, and M. musculus. All these observations suggest that Spo11 is directly involved in catalyzing DSB formation to trigger meiotic recombination. Figure \(10\) provides an overview of this process. Panel (A) shows a schematic of the formation of haploid gametes from a diploid cell with a single pair of homologous chromosomes. DSB formation and recombination promote homolog pairing and lead to the exchange of chromosomal fragments (crossovers) in the context of synapsed chromosomes. Panel (B) shows meiotic recombination is initiated by Spo11-mediated DSB formation and leads to the formation of crossovers via a ZMM-dependent double Holliday Junction (dHJ) resolution pathway or non-crossovers by synthesis-dependent strand annealing. Panel (C) shows the relationships between meiotic recombination and higher-order chromosome structure. DSB formation happens in the context of the loop-axis structure. As recombination progresses, the SC polymerizes between the axes and is disassembled before chromosome segregation. Axis proteins Red1 (red ovals) and Hop1 (yellow ovals) are shown. Panel (D)shows that in metaphase I, homologs are held together through chiasmata and sister chromatid cohesion. Image from: Following break formation, Spo11 remains covalently attached to the 5′-strands at both DNA ends and is released by an endonucleolytic cleavage reaction mediated by MRX (Mre11, Rad50, and Xrs2) and Sae2, which liberates Spo11 attached to a short oligonucleotide (Fig. 25.3.7B). The 5′-strands are further resected by 5′-3′ exonucleases to produce long single-stranded tails, which are coated with the ssDNA-binding protein, RPA. RPA is then replaced by recombinases Rad51 and Dmc1 that form a nucleoprotein filament and search for sequence similarity preferentially located on the homologous chromosome, producing D-loop structures. Following DNA synthesis using the homolog as a repair template, the recombination structures experience one of two main outcomes (Fig. 25.3.7B). The invading strand can be ejected from the donor by the action of helicases, which provides an opportunity for the DNA ends to re-anneal. This process is referred to as synthesis-dependent strand annealing (SDSA) and produces non-crossovers, that is, products not associated with reciprocal exchanges of chromosome fragments, but with the local transfer of genetic information from the repair template to the broken molecule (gene conversion). Alternatively, recombination structures are stabilized by the “ZMM” family of proteins and channeled through a pathway that produces mostly crossovers. Here, both ends of the break engage the donor to form a double Holliday Junction intermediate, which is resolved through a crossover-specific pathway that involves MutLγ and Exo1. Every aspect of meiotic recombination is tied to the structural organization of the chromosomes (Fig. 25.3.7C). Early in the meiotic prophase, chromosomes organize as a series of DNA loops that are anchored along a nucleoprotein axis. DSB formation happens in the context of this loop-axis structure. As recombination progresses, polymerization of a proteinaceous structure called the synaptonemal complex (SC) initiates between the two axes and elongates along their entire length. Recombination proceeds within the SC, inside a nodule embedded between the axes. After recombination is completed, the SC disassembles and crossovers, now cytologically visible as chiasmata, provide physical connections between the homologs until their segregation at anaphase (Fig. 25.3.7D).

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/24%3A_DNA_Metabolism/24.03%3A_DNA_Recombination.txt

Search Fundamentals of Biochemistry Types of RNA Structurally speaking, ribonucleic acid (RNA), is quite similar to DNA. However, whereas DNA molecules are typically long and double-stranded, RNA molecules are much shorter and are typically single-stranded. A ribonucleotide within the RNA chain contains ribose (the pentose sugar), one of the four nitrogenous bases (A, U, G, and C), and a phosphate group. The subtle structural difference between the sugars gives DNA added stability, making DNA more suitable for the storage of genetic information, whereas the relative instability of RNA makes it more suitable for its more short-term functions. The RNA-specific pyrimidine uracil forms a complementary base pair with adenine and is used instead of the thymine that is found in DNA. Even though RNA is single-stranded, most types of RNA molecules show extensive intramolecular base pairing between complementary sequences within the RNA strand, creating a predictable three-dimensional structure essential for their function, as shown in Figure $1$ and Figure $2$. RNA can largely be divided into two types, one that carries the code for making proteins or coding RNA, which is also called messenger RNA (mRNA), and non-coding RNA (ncRNA). The ncRNA can be subdivided into several different types, depending either on the length of the RNA or on the function. Size classification begins with the short ncRNAs (~20–30 nt), which include microRNAs (miRs), and small interfering (siRNAs); the small ncRNAs up to 200 nt, which include transfer RNA (tRNA), small nuclear RNA (snRNA), and small nucleolar RNA (snoRNA); and long ncRNAs ( > 200 nt), which include ribosomal RNA (rRNA), enhancer RNA (eRNA) and long intergenic ncRNAs (lincRNAs), among others. Cells access the information stored in DNA by creating RNA, through the process of transcription, which then directs the synthesis of proteins through the process of translation. The three main types of RNA directly involved in protein synthesis are messenger RNA (mRNA), ribosomal RNA (rRNA), and transfer RNA (tRNA). The mRNA carries the message from the DNA, which controls all of the cellular activities in a cell. If a cell requires a certain protein to be synthesized, the gene for this product is “turned on” and the mRNA is synthesized through the process of transcription. The mRNA then interacts with ribosomes and other cellular machinery to direct the synthesis of the protein it encodes during the process of translation. mRNA is relatively unstable and short-lived in the cell, especially in prokaryotic cells, ensuring that proteins are only made when needed. rRNA and tRNA are stable types of RNA. In prokaryotes and eukaryotes, tRNA and rRNA are encoded by the DNA, where they are transcribed into long RNA molecules that are subsequently cut to release smaller fragments containing the individual mature RNA species. In eukaryotes, synthesis, cutting, and assembly of rRNA into ribosomes takes place in the nucleolus region of the nucleus, but these activities occur in the cytoplasm of prokaryotes. Within the nucleolus region, ribosome assembly requires the activity of numerous snoRNAs. Ribosomes are composed of rRNA and protein. As its name suggests, rRNA is a major constituent of ribosomes, composing up to about 60% of the ribosome by mass and providing the location where the mRNA binds. The rRNA ensures the proper alignment of the mRNA, tRNA, and the ribosomes; the rRNA of the ribosome also has an enzymatic activity (peptidyl transferase) and catalyzes the formation of the peptide bonds between two aligned amino acids during protein synthesis (Figure 26.1.3). Although rRNA had long been thought to serve primarily a structural role, its catalytic role within the ribosome was shown in 2000. Scientists in the laboratories of Thomas Steitz (1940–) and Peter Moore (1939–) at Yale University were able to crystallize the ribosome structure from Haloarcula marismortui, a halophilic archaeon isolated from the Dead Sea. Because of the importance of this work, Steitz shared the 2009 Nobel Prize in Chemistry with other scientists who made significant contributions to the understanding of ribosome structure. The structure and function of ribosomes will be discussed in further detail in Chapter 27. Transfer RNA (tRNA) is the third prominent type of RNA involved in protein translation. tRNAs are usually only 70–90 nucleotides long. They carry the correct amino acid to the site of protein synthesis in the ribosome. It is the base pairing between the tRNA and mRNA that allows for the correct amino acid to be inserted in the polypeptide chain being synthesized, as shown in Figure $3$. Any mutations in the tRNA or rRNA can result in global problems for the cell because both are necessary for proper protein synthesis. As described previously, some RNA molecules have enzymatic properties and serve as ribozymes. Within this chapter, the activity of snRNAs during the process of intron removal from mRNA sequences function as ribozymes and will be described. Furthermore, a detailed description of the enzymatic features of the ribosome structure will be provided in Chapter 27. Other small ncRNAs and lncRNA molecules play a role in the regulation of transcriptional and translational processes. For example, the post-transcriptional expression levels of many genes can be controlled by RNA interference, in which miRNAs, specific short RNA molecules, pair with mRNA regions and target them for degradation, as shown in Figure $4$. This process is aided by protein chaperones called argonautes. This antisense-based process involves steps that first process the miRNA so that it can base-pair with a region of its target mRNAs. Once the base pairing occurs, other proteins direct the mRNA to be destroyed by nucleases. Fire and Mello were awarded the 2006 Nobel Prize in Physiology or Medicine for this discovery. At steady state, the vast majority of human cellular RNA consists of rRNA (∼90% of total RNA for most cells) as shown in Figure $5$. Although there is less tRNA by mass, their small size results in their molar level being higher than rRNA (Figure 26.1.5). Other abundant RNAs, such as mRNA, snRNA, and snoRNAs are present in aggregate at levels that are about 1–2 orders of magnitude lower than rRNA and tRNA (Figure 26.1.5). Certain small RNAs, such as miRNA and piRNAs can be present at very high levels; however, this appears to be cell type dependent. lncRNAs are present at levels that are two orders of magnitude less than total mRNA. Although the estimated number of different types of human lncRNAs may have a very restricted expression pattern and thus, accumulate to higher levels within specific cell types. For example, sequencing of mammalian transcriptomes has revealed more than 100,000 different lncRNA molecules can be produced, compared with the approximate 20,000 protein-coding genes. The diversity and functions of the transcriptome within biological processes are currently a highly active area of research. RNA Polymerases This chapter will focus on the synthesis of RNA by DNA-dependent RNA Polymerase Enzymes (RNAPs). These enzymes are required to carry out the process of transcription and are found in all cells ranging from bacteria to humans. All RNAPs are multi-subunit assemblies, with bacteria having five core subunits, α2ββ'ω, that have homologs in archaeal and eukaryotic RNAPs. Bacterial RNAPs are the simplest form of RNA polymerases and provide an excellent system to study how they control transcription. Prokaryotic RNA Polymerase Enzymes The RNAP catalytic core within bacteria contains five major subunits (α2ββ'ω) (see Figure $7$) below. To position this catalytic core onto the correct promoter requires the association of a sixth subunit called the sigma factor (σ). Within bacteria, there are multiple different sigma factors that can associate with the catalytic core of RNAP that help to direct the catalytic core to the correct DNA locations, where RNAP can then initiate transcription. For example, within E. coli σ70 is the housekeeping sigma factor that is responsible for transcribing most genes in growing cells. It keeps essential genes and pathways operating. Other sigma factors are activated during certain environmental situations, such as σ38 which is activated during starvation or when cells reach the stationary phase. When the sigma subunit associates with the RNAP catalytic core, the RNAP has then formed the holoenzyme. When bound to DNA, the holoenzyme conformation of RNAP can initiate transcription. Transcription takes place in several stages. To start with, the RNA polymerase holoenzyme locates and binds to promoter DNA. At this stage the RNAP holoenzyme is it the closed conformation (RPc), as shown in Figure $6$. Initial specific binding to the promoter by sigma factors of the holoenzyme sets in motion conformational changes in which the RNAP molecular machine bends and wraps the DNA with mobile regions of RNAP playing key roles, as shown in Figure $6$. Next, RNAP separates the two strands of DNA and exposes a portion of the template strand. At this point, the DNA and the holoenzyme are said to be in an ‘open promoter complex’ (RPo), and the section of promoter DNA that is within it is known as a ‘transcription bubble’. Intermediates(I1-3) between RPc and RPo occur. In bacterial systems, the sigma factor locates the transcriptional start site using key DNA sequence elements located at -35 nucleotides and -10 nucleotides from the transcriptional initiation site, as shown in Panel A of Figure $7$. This region is called the Pribnow box. For RNAP from Thermus aquaticus, the −35 element interacts exclusively with σ4. The duplex DNA just upstream of the −10 element (−17 to −13) interacts with β′, σ2, and σ3 (Panel B). Flipping of the A−11(nt) base from the duplex DNA into its recognition pocket in σ2 is thought to be the key event in the initiation of promoter melting and the formation of the transcription bubble (Panel C). Once the transcription bubble has formed and transcription initiates, the sigma subunits dissociate from the complex and the RNAP catalytic subunit continues elongation on its own. Panel (A) shows oligonucleotides used for the crystallization of the RNAP holoenzyme in the open conformation. The numbers above denote the DNA position with respect to the transcription start site (+1). The −35 and −10 (Pribnow box) elements are shaded yellow, and the extended −10 and discriminator elements are purple. The nontemplate-strand DNA (top strand) is colored dark grey; template-strand DNA (bottom strand), light grey; RNA transcript, red. Panel (B) shows the overall structure of RNAP holoenzyme in the open conformation bound with the DNA nucleotides. The nucleic acids are shown as CPK spheres and color-coded as in diagram A. Within RNAP, the αI, αII, ω, are shown in grey; β in light cyan; β′ in light pink; Δ1.1σA in light orange. The Taq EΔ1.1σA (Taq derives from Thermus aquaticus) is shown as a molecular surface and the forward portion of the RNAP holoenzyme is transparent to reveal the RNAP active site Mg2+ (yellow sphere) and the nucleic acids held inside the RNAP active site channel. Panel (C) Electron density and model for RNAP holoenzyme nucleic acids in the open conformation. Color coding matches diagram A. Figure $8$ shows an interactive iCn3D model of the T. aquaticus transcription initiation complex containing bubble promoter and RNA (4XLN). It is colored coded in fashion similar to that shown in Figure $7$. The rendering of the DNA is as follows: • The non-template (nt-strand) DNA is colored dark grey; spheres • template (t-strand) DNA, light grey; spheres • Pribnow box yellow • discriminator purple The proteins are shown as surfaces with transparent secondary structures underneath. The color is as follows: • (αI, αII, ω, light yellow; • β, light cyan; • β′, light pink; • Taq EΔ1.1σA, light orange), The RNA polymerase active site is located at the Mg2+ (black sphere) binding site. The nucleic acids are inside the RNAP active site channel. Eukaryotic RNA Polymerase Enzymes In eukaryotic cells, three RNAPs (I, II, and III) share the task of transcription, the first step in gene expression. RNA Polymerase I (Pol I) is responsible for the synthesis of the majority of rRNA transcripts, whereas RNA Polymerase III (Pol III) produces short, structured RNAs such as tRNAs and 5S rRNA. RNA Polymerase II (Pol II) produces all mRNAs and most regulatory and untranslated RNAs. The three eukaryotic RNA polymerases contain homologs to the five core subunits found in prokaryotic RNAPs. In addition, the eukaryotic Pol I, Pol II and Pol III have five additional subunits forming a catalytic core that contains 10 subunits, as shown in Figure $9$. The core has a characteristic crab-claw shape, which encloses a central cleft that harbors the DNA, and has two channels, one for the substrate NTPs and the other for the RNA product. Two ‘pinchers’, called the ‘clamp’ and ‘jaw’ stabilize the DNA at the downstream end and allow the opening and closing of the cleft. For transcription to occur, the enzyme has to maintain a transcription bubble with separated DNA strands, facilitate the addition of nucleotides, translocate along the template, stabilize the DNA:RNA hybrid, and finally allow the DNA strands to reanneal. This is achieved by a number of conserved elements in the active site, which include the fork loop(s), rudder, wall, trigger loop, and bridge helix. DNA (black) is melting into a transcription bubble that allows template-strand pairing with RNA (red) in a 9-10 base pair RNA-DNA hybrid. The bridge helix (cyan) and trigger loop/helices (yellow/orange) lie on the downstream side of the active site. The presumed path of the NTP entry is indicated by the straight arrow. Interconversion of the trigger loop and trigger helices is indicated by the curved arrow. The RNA polymerase subunits are shown as semi-transparent surfaces with the identities of orthologous subunits in bacteria (α, β, and β', gray, blue, and pink, respectively), archaea (D, L, B, and A), and eukaryotic RNA polymerase II (RPB3, 11, RPB2, RPB1) indicated. The active site Mg2+ ions are shown as yellow spheres, and α,β-methylene-ATP in green and red. Table $1$ shows RNA polymerase (RNA pol) subunit composition in bacteria, archaea, and, yeast and plants (both eukaryotes). Bacteria Archaea RNA pol I RNA pol II RNA pol III RNA pol IV (plants) RNA pol V (plants) β Rpo1 (RpoA) RPA190 RPB1 RPC160 NRPD1 NRPE1 β' Rpo2 (RpoB) RPBA135 RPB2 RPC128 NRPD/E2 NRPD/E2 α Rpo3 (RpoD) RPAC40 RPB3 RPAC40 RPB3 [1] RPB3 [1] α Rpo11 (RpoL) RPAC19 RPB11 RPAC19 RPB11 RPB11 ω Rpo6 (RpoK) RPB6 RPB6 RPB6 RPB6 [1] RPB6 Rpo5 (RpoH) RPB5 RPB5 RPB5 RPB5 [3] NRPES5 Rpb8 (RpoG)* RPB8 RPB8 RPB8 RPB8 [1] RPB8 [1] Rpo10 (RpoN) RPB10 RPB10 RPB10 RPB10 RPB10 Rpo12 (RpoP) RPB12 RPB12 RPB12 RPB12 RPB12 Rpo4 (RpoF) RPA14 RPB4 RPC17 NRPD/E4 NRPD/E4 Rpo7(RpoE) RPA43 RPB7 RPC25 NRPD7 [1] NRPE7 RPA12 RPB9 RPC11 NRPD9b RPB9 Rpo13* RPA49   RPC53 RPA34.5   RPC37 RPC82 RPC34 RPC31 Table $1$: RNA polymerase (RNA pol) subunit composition. Abel, C., Verónica, M., I., G. A. , & Francisco, N. (2017). Subunits Common to RNA Polymerases. In (Ed.), The Yeast Role in Medical Applications. IntechOpen. https://doi.org/10.5772/intechopen.70936. Creative Commons Attribution 3.0 License Schematic representations of the structure of the eukaryotic RNA pols I, II and III are shown in Figure $10$. Figure $10$: Schematic representation of the structure of the RNA pols I, II and III. Each RNA pol common subunit is indicated in grey. The numbers correspond to each subunit are indicated in Subunits Common to RNA Polymerases. Abel et al, ibid. RNA polymerases must bind to DNA, and to host of transcription factors (TF) necessary for specific and regulatable transcription. (Note: RNAP is not considered a transcription factor.) The comparative structures of RNAP I-III are shown in Figure $11$. The "stalk" is a structural feature found in eukaryotes but not in prokaryotes. The figure focuses mostly on a comparison of RNAP I and RNAP III. Figure $11$: Comparison of RNAPI, II and III structures and transcription factors. Turowski TW and Boguta M (2021) Specific Features of RNA Polymerases I and III: Structure and Assembly. Front. Mol. Biosci. 8:680090. doi: 10.3389/fmolb.2021.680090. Creative Commons Attribution License (CC BY). Panel (A) shows the general architecture of RNAPII, consisting of the catalytic core and stalk. RNAPII core consists of a DNA binding channel, catalytic center, and assembly platform. RNAPII binds multiple transcription factors (TFs). Some TFs are homologous to additional subunits of specialized RNAPs (i.e., TFIIF). Panel (B) shows the subunit composition of eukaryotic RNAPs. Human nomenclature is shown for comparison. Please note that the C-terminal region of Rpa49 subunit harbors a “tandem winged helix” which is predicted in TFIIE and that human RNAPIII RPC7 subunit is coded by two isoforms α and β. The question mark indicates the name is unconfirmed. Panel (C) shows the subunit composition of yeast RNAPI. Panel (D) shows a Model of the RNAPI pre-initiation complex, showing an early intermediate with visible Rrn3 and core factor (CF). TATA-binding protein (TBP) and upstream-associated factor (UAF) are added schematically. Panel (E) shows the subunit composition of yeast RNAPIII. Panle (F) shows atomic model of RNAPIII pre-initiation complex with TFIIIB. The Rpc82/34/31 heterotrimer is involved in initiation and marked in green as in E. TFIIIC is added schematically. PDB: 5C4X, 5FJ8, 4C3J, 6EU0, and 6TPS The stalks have two proteins that are not as homologous as the core subunits. These are highlighted in blue in Panel B of Figure $11$. RNAP I and III have additional subunits compared to RNAP II. Overall RNAP I-III have 14, 12, and 17 subunits, respectively. RNAP I and III appear to have integrated transcription-like factors into their core enzyme. In contrast, RNAP II (which transcribes DNA to form messenger RNA), binds to discrete and separate transcription factors to form a preinitiation complex (PIC)which we will discuss below. Transcription Factors and the Preinitiation Complex (PIC) Unlike prokaryotic systems which can initiate the recruitment of RNAP holoenzymes directly onto the DNA promoter regions and mediate the conversion of RNAP to the open conformation, eukaryotic RNA polymerases require a host of additional general transcription factors (GTFs), to enable this process. Here we will focus on the activation of RNA Polymerase II as an example of the complexity of eukaryotic transcription initiation. Class II gene transcription in eukaryotes is a tightly regulated, essential process controlled by a highly complex multicomponent machinery. A plethora of proteins, more than a hundred in humans, are organized in very large multiprotein assemblies that include a core of General Transcription Factors (GTFs). The GTFs include the factors TFIIA, TFIIB, TFIID, TFIIE, TFIIF, TFIIH, RNA polymerase (RNA pol II), as well as a large number of diverse complexes that act as co-activators, co-repressors, chromatin modifiers, and remodelers, as shown in Figure $12$. Class II gene transcription is regulated at various levels: while assembling on chromatin, before and during transcription initiation, throughout elongation and mRNA processing, and termination. A host of activators and repressors has been reported to regulate transcription, including a central multisubunit complex called the Mediator that helps in the recruitment of GTFs and the activation of RNA Pol II. Here we will focus on the formation of the GTFs that make up the core preinitiation complex (PIC) during transcriptional activation. Class II gene transcription in humans is brought about by over a hundred polypeptides assembling on the core promoter of protein-encoding genes, which then give rise to mRNA. A PIC on a core promoter is shown in a schematic representation. PIC contains, in addition to promoter DNA, the GTFs (TFIIA, B, D, E, F, and H), and RNA Pol II. PIC assembly is thought to occur in a highly regulated, stepwise fashion, as indicated. TFIID is among the first GTFs to bind the core promoter via its TATA-box Binding Protein (TBP) subunit. Nucleosomes at transcription start sites contribute to PIC assembly, mediated by signaling through epigenetic marks on histone tails. The Mediator (not shown) is a further central multiprotein complex identified as a global transcriptional regulator. TATA = TATA-box DNA; BREu = B recognition element upstream; BREd = B recognition element downstream; Inr = Initiator; DPE = Down-stream promoter element. Figure from: Transcription of RNA pol II-dependent genes is triggered by the regulated assembly of the Preinitiation Complex (PIC). PIC formation starts with the binding of TFIID to the core promoter. TFIID is a large megadalton-sized multiprotein complex with around 20 subunits made up of 14 different polypeptides: the TATA-box binding protein (TBP) and the TBP-associated factors (TAFs) (numbered 1–13), as shown in Figure $13$. Some of the TAF subunits are present in two copies. A key feature in TAFs is the histone fold domain (HFD), which is present in 9 out of 13 TAFs in TFIID. The HFD is a strong protein–protein interaction motif that mediates specific dimerization. The HFD-containing TAFs are organized in discrete heterodimers, with the exception of TAF10, which is capable of forming dimers with two different TFIID components, TAF3 and TAF8. HFDs and several other structural features of TBP and the TAFs are well conserved between species. TFIID is a large megadalton-sized multiprotein complex comprising about 20 subunits made up of 14 different polypeptides. The constituent proteins of TFIID, TBP and the TAFs, are shown in a schematic representation depicted as bars (inset, left). Structured domains are marked and annotated. The presumed stoichiometry of TAFs and TBP in the TFIID holo-complex is given (far left, gray underlaid). TAF10 (in italics) makes histone fold pair separately with both TAF3 and TAF8. TAFs present in a physiological TFIID core complex extracted from eukaryotic nuclei are labeled in bold. The architecture of the TFIID core complex (EMD-2230) determined by cryo-EM is shown (bottom left) in two views related by a 90° rotation (arrows). The holo–TFIID complex is characterized by remarkable structural plasticity. Two conformations, based on cryo-EM data (EMD-2284 and EMD-2287), are shown on the right, a canonical form (top) and a more recently observed rearranged form (bottom). In the rearranged conformation, lobe A (colored in red) migrates from one extreme end of the TFIID complex (attached to lobe C) all the way to the other extremity (attached to lobe B). TFIID was shown to adopt an asymmetric, horse-shoe shape with three almost equal-sized lobes (A, B, and C), exhibiting a considerable degree of conformational flexibility with at least two distinct conformations (open and closed), as shown in Figure $12$. The TBP component of TFIID binds with a specific DNA sequence called the TATA box. This DNA sequence is found around 30 base pairs upstream of the transcription start site in many eukaryotic gene promoters. When TBP binds to a TATA box within the DNA, it distorts the DNA by inserting amino acid side chains between base pairs, partially unwinding the helix, and doubly kinking it. The distortion is accomplished through a great amount of surface contact between the protein and DNA. TBP binds with the negatively charged phosphates in the DNA backbone through positively charged lysine and arginine amino acid residues. The sharp bend in the DNA is produced through the projection of four bulky phenylalanine residues into the minor groove. As the DNA bends, its contact with TBP increases, thus enhancing the DNA-protein interaction. The strain imposed on the DNA through this interaction initiates the melting, or separation, of the strands. Because this region of DNA is rich in adenine and thymine residues, which base-pair through only two hydrogen bonds, the DNA strands are more easily separated. The role of TAFs is complicated. Take for example TAF11 and TAF13. These act as competitive inhibitors of TBP to the TATA (Pribnow) box as well as TAF1 which somewhat mimics the structural features of the Pribnow box. TAF11/TAF13 binds to the DNA surface where TBP binds. suggesting a novel regulation of TFIID. These interactions are illustrated in Figure $14$. Figure $14$: Novel TFIID regulatory state comprising TAF11/TAF13/TBP. Kapil Gupta,et al. (2017) Architecture of TAF11/TAF13/TBP complex suggests novel regulation properties of general transcription factor TFIID eLife 6:e30395. https://doi.org/10.7554/eLife.30395. Creative Commons Attribution License Given its role in transcribing the DNA for thousands of messenger RNAs, let's focus on the preinitation complex for RNAP II. Structures are known for the closed and open promoters. A key component is TFH (see Figure $12$). TFIIH opens the DNA for transcription. As if we need to complicate the structure of the preinitiation complex even more, it turns out the TFIIH is not a single dark green cartoon as shown in Figure 12, but a rather large complex itself. Its structure is shown in Figure $15$. Figure $15$: Structure of the TFIIH core complex. Greber et al. (2019). The complete structure of the human TFIIH core complex. eLife 8:e44771. https://doi.org/10.7554/eLife.44771. Creative Commons Attribution License Panes (A, B, C) shows three views of the structure of the TFIIH core complex and MAT1. Subunits are color-coded and labeled (in color); individual domains are labeled (in black) and circled if needed for clarity. Panel (D) shows the domain-level protein-protein interaction network between the components of the TFIIH core complex and MAT1 derived from the interactions observed in our structure. Proteins are shown with the same colors as in A and major unmodeled regions are shown in grey. Abbreviations: CTD: C-terminal domain; DRD: DNA damage recognition domain; FeS: iron sulfur cluster domain; NTD: N-terminal domain; vWFA: von Willebrand Factor A. The largest subunits are DNA-dependent helicases/translocases/ATPases XPB and XPD, which are bridged by MAT1. It appears that XPB starts and propagates a twist in the DNA which propagates ot open the DNA 30 BP downstream of the TATA box. This mostly likely is followed by the dissociation of TFIIH and a stoppage in DNA twisting, which allows RNA transcription to start. Figure $16$ shows an interactive iCn3D model of the human/mouse/mastadenovirus C RNA polymerase II core pre-initiation complex with open promoter DNA (7NVU). The following color schemes are used: • MAT1 - orange • TF2A - yellow • TF2B - green • TF2E - magenta • TF2F- purple • ATP-dependent translocase (helicase) subunit XPD - dark slate gray • TBP (TATA box binding protein) - red • TF2H - Pink • 2H-XPB helicase - maroon • NT-DNA - cyan • template (T)-DNA - blue (Note: part of it is shown in a yellow cartoon in the interactive model) • the SF4 cofactor is shown in spacefill with CPK colors In summary, the binding of TFIID to the core promoter is followed by the recruitment of further GTFs and RNA pol II. Several lines of evidence suggest that this process occurs in a defined, stepwise order and undergoes significant restructuring. First, PIC adopts an inactive state, the “closed” complex, which is incompetent to initiate transcription. In addition to TFIID, TFIIH is also critical for the shift of RNA Pol II from the closed to the open conformation. TFIIH has an ATP-dependent translocase activity within one of its subunits, that opens up about 11 to 15 base pairs around the transcription start site by moving along one DNA strand inducing torsional strain, leading to conformational rearrangements and the positioning of single-stranded DNA to the active site of RNA pol II. In this “open” complex, RNA pol II can enter elongation to transcribe throughout a gene in a highly processive manner without dissociating from the DNA template or losing the nascent RNA. In most eukaryotes, after synthesizing about 20–100 bases, RNA pol II can pause (Promoter proximal pause) and then disconnect from promoter elements and other components of the transcription machinery, giving rise to a fully functional elongation complex in a process called promoter escape. The promoter-bound components of the PIC, in contrast, remain in place, and thus only TFIIB, TFIIF, and RNA pol II need to be recruited for re-initiation, significantly increasing the transcription rate in subsequent rounds of transcription. Promoter escape is preceded by an abortive transcription in many systems, where multiple short RNA products of 3 to 10 bases in length are synthesized. In addition to promoter elements within the DNA, enhancer elements are also important for the initiation of transcription. Promoters are defined as DNA elements that recruit transcription complexes for the synthesis of coding and non-coding RNA. Enhancers are defined as DNA elements that positively regulate transcription at promoters over long distances in a position- and orientation-independent manner. However, studies have revealed that many enhancers can recruit Pol II and initiate transcription of enhancer RNA (eRNA), thus blurring the functional distinction between enhancers and promoters (Figure 10.13). Enhancer transcription produces relatively short ncRNA. Furthermore, transcription at enhancers is unstable and often leads to the termination of elongation. In contrast, transcription initiation at most Pol II promoters is stable and produces long mRNAs. Topological studies revealed that enhancers come in close proximity to target gene promoters during transcription activation. According to current gene activation models, the Mediator complex forms a physical bridge between distant regulatory regions and promoters, thereby promoting looping. Transcription of at least a subset of genes regulated by enhancers occurs in bursts indicating a discontinuous process of transcription complex recruitment, assembly, and/or conversion to elongation-competent forms. The bursting phenomenon suggests that enhancer/promoter contacts may be transient and infrequent, as shown in Figure $17$. Depicted are the steps involved in the recruitment of Pol II to SEs, assembly into elongation-competent transcription complexes, transcription initiation, and elongation, abortion and termination, and transfer to target genes. Transcription factors recruit Mediator and other co-regulators to SEs. Mediator recruits Pol II and assembles a fraction into elongation competent transcription complexes. Transcription is initiated by phosphorylation of the CTD. Early abortion and transcription termination conferred by Integrator releases Pol II, which is dephosphorylated and transferred to target gene promoters. Super Enhancer Element (SE). Transcriptional Elongation and Termination Prokaryotic Transcriptional Elongation The rate of transcription elongation by E. coli RNAP is not uniform. RNA synthesis is characterized by pauses, some of which may be brief and resolved spontaneously, whereas others may lead to the transcription elongation complex (TEC) backtracking. Elongation rate and pausing are determined by template sequence and RNA structure (e.g., stem-loops) and involve at least two components of the RNAP catalytic center, the bridge helix (BH) and trigger loop (TL). Elongation is proposed to occur in three steps, as shown in Figure $18$. First, the TL folds in response to NTP binding. Mutational analyses indicate that this conformational change in the TL can be rate-limiting, and reflects the ability of the incoming NTP to bind to TEC. The second step is the incorporation of the NTP and the release of pyrophosphate. The third step involves the translocation of the RNAP down the DNA Template such that the next RNA nucleotide can be added to the nascent transcript. The trigger loop hinges, bridge helix hinges, and bridge helix bending models are based on molecular dynamics simulations. At the top of the figure, diagrams of the closed TEC, the closed product TEC (after chemistry), and the translocating TEC are shown. DNA is grey; RNA is red; the NTP substrate (or incorporated NMP and pyrophosphate) is blue; the trigger loop (TL) is purple; the bridge helix (BH) is yellow. Interpretations of simulations are shown schematically below. Simulations indicate trigger loop hinges H1 and H2, bridge helix hinges H3 and H4 and bridge helix bend modes B1 (straighter) and B2 (more sharply bent). Backtracking of TEC may take place after a brief pause in transcription, caused by the thermodynamic properties of nucleic acids sequences surrounding the elongation complex. In addition,  misincorporation events render elongation complexes prone to backtracking by at least one bp. In this case, the rescue from backtracking through the cleavage of the 3' end of the erroneous transcript also may be seen as a proofreading reaction. Any backtracking event causes a pause or arrest of transcription elongation, which may limit its overall rate (the average speed of RNAP along the template) or the processivity (the fraction of RNAP molecules reaching the end of the gene). While the general structure of the elongation complex (the transcription bubble, the RNA-DNA hybrid) remains unchanged during backtracking, the extension of RNA becomes impossible in this conformation. However, such complexes can be resolved by the hydrolytic activity of RNAP, which cleaves the phosphodiester bond in the active center of the backtracked complex, producing a new RNA 3' end in the active center. For single base backups, the hydrolytic reaction is catalyzed by a flexible domain of RNAP located in the secondary channel called the Trigger Loop (TL) and the two metal ions of the active center. Longer sequences of backtracked TEC can restart when acted upon by GreA/B factors, which restore the 3'-end of the nascent transcript to the active center. GreA and GreB are transcript cleavage factors that act on backtracked elongation complexes. When Gre factors are bound in the secondary channel, Gre factors displace the TL from the active center, as shown in Figure $19$. The displacement switches off the relatively slow TL-dependent intrinsic transcript hydrolysis, and imposes the highly efficient Gre-assisted hydrolysis. This efficiency is thought to be due to the stabilization of the second catalytic Mg2+ ion and an attacking water molecule by the Gre factors. Panel (A) shows a ribbon diagram of the GreA and GreB proteins. Panel (B) shows the mode of functioning of Gre factors. The Gre factor is bound to the active elongation complex but does not impose hydrolytic activity on it. Upon backtracking or misincorporation, the Gre factor protrudes its coiled-coil domain through the secondary channel of RNAP (shown in the lefthand diagram), where it substitutes for the catalytic domain Trigger Loop (TL). This substitution switches off the slow TL-dependent phosphodiester bond hydrolysis and, and instead, facilitates highly efficient Gre-dependent hydrolysis. After the resolution of the backtracked complex through RNA cleavage, the elongation complex returns to the active conformation, and the Gre factor gives way to the TL, which can now continue the catalysis of RNA synthesis (shown in the right-hand diagram). The controlled switching between Gre and the TL eliminates possible interference of Gre with the RNA synthesis. Prokaryotic Transcriptional Termination Transcription termination determines the ends of transcriptional units by disassembling the transcription elongation complex (TEC), thereby releasing RNA polymerases and nascent transcripts from DNA templates. Failure in termination causes transcription readthrough, which yields wasteful and possibly harmful intergenic transcripts. It can also perturb the expression of downstream genes when the unterminated TEC sweeps transcription initiation complexes off their promoters or collides with RNA polymerases that transcribe opposite strands. Transcriptional termination in prokaryotes can be template-encoded and factor-independent (intrinsic termination), or require accessory factors, such as Rho, Mfd, and DksA. Intrinsic termination occurs at specific template sequences - an inverted repeat followed by a run of A residues. Termination is driven by the formation of a short stem-loop structure in the nascent RNA chain, as shown in Figure $20$. RNA synthesis arrests and TEC dissociates at the 7th and 8th U of the run. Formation of the stem-loop dissociates the weak rU:dA hybrid. Stem-loop formation is hindered by upstream complementary RNA sequences that compete with the downstream portion of the stem, as well as by RNA: protein interactions in the RNA exit channel. Intrinsic termination depends critically upon timing. Hairpin folding and transcription of the termination point must be coordinated, so that the complete hairpin is formed by the time RNAP transcribes the termination point. The size of the stem, the sequence of the stem, and the length of the loop all affect termination efficiency. The bridge α-helix in the β' subunit borders the active site and may have roles in both catalysis and translocation. Mutations in the YFI motif (β' 772-YFI-774) affect intrinsic termination as well as pausing, fidelity, and translocation of RNAP. One mutation, F773V, abolishes the activity of the λ tR2 intrinsic terminator, although neighboring mutations have little effect on termination. Modeling suggests that this unique phenotype reflects the ability of F773 to interact with the fork domain in the β subunit. Panel (A) shows the open conformation of the RNAP during transcriptional elongation. RNAP is shown in yellow, the DNA template in blue, and the nascent RNA in red. Key elements of the RNAP RNA exit channel are shown in grey and labeled as indicated. Panel (B) shows the extension of the nascent RNA through the RNAP exit channel and the potential for forming the RNA hairpin structure when enough length has been achieved. Panel (C) shows the clamp opening and disintegration of the TEC when the RNA hairpin structure is encountered at the transcriptional bubble. Figure $21$ shows an interactive iCn3D model of the T. thermophilus RNAP polymerase elongation complex with the NTP substrate analog (2O5J). (long load time) Transcriptional termination can also be dependent upon accessory factors, such as the Rho protein. Transcription termination factor Rho is an essential protein in E. coli first identified for its role in transcription termination at Rho-dependent terminators, and is estimated to terminate ~20% of E. coli transcripts. The rho gene is highly conserved and nearly ubiquitous in bacteria. Rho is an RNA-dependent ATPase with RNA:DNA helicase activity, and consists of a hexamer of six identical monomers arranged in an open circle, as shown in Figure $22$. Rho binds to single-stranded RNA in a complex multi-step pathway that involves two distinct sites on the hexamer. The primary binding site (PBS), distributed on the N-terminal domains around the hexamer (cyan), ensures initial anchoring of Rho to the transcript at a Rut (Rho utilization) site, a∼70 nucleotides (nt) long, cytidine-rich and poorly-structured RNA sequence. Each Rho monomer contains a subsite capable of binding specifically the base residues of a 5′-YC dimer (Y being a pyrimidine). Biochemical and structural data suggest that Rho initially binds to RNA in an open, ‘lock-washer’ conformation that closes into a planar ring as RNA transfers to the central cavity. There, the ssRNA contacts an asymmetric secondary binding site (SBS) (green), and this step, which presumably is rate-limiting for the overall reaction, leads to motor activation. Upon hydrolysis of ATP, the ssRNA is pulled upon conformational changes of the conserved Q and R loops of the SBS, leading to Rho translocation, and ultimately promoting RNA polymerase (RNAP) dissociation. The molecular mechanism of Rho translocation based on single-molecule fluorescence methods appears to be tethered tracking. The tethered tracking model postulates that Rho maintains its contacts between the PBS and the loading (Rut) site upon translocation (Panel B). This mechanism would allow Rho to maintain its high-affinity interaction with Rut, and implies the growth of an RNA loop between the PBS and the SBS upon translocation. Panel (A) shows the molecular structure of the Rho protein (PDB 1pv4) Panel (B)shows how Rho assembles as a homo-hexameric ring (red spheres or tetragons), with RNA (black/yellow curve) binding to the primary binding sites (PBS, cyan) and the secondary binding sites inside the ring (SBS, green), where ATP-coupled translocation takes place. The Rut-specific binding site is depicted in yellow. The tethered-tracking model proposed that Rho translocates RNA while maintaining interactions between PBS and Rut. This model requires the formation of a loop that would shorten the extension of RNA upon translocation. Figure modified from: Figure $23$ shows an interactive iCn3D model of the E. Coli Rho transcription termination factor in complex with ssRNA substrate and ANPPNP (1PVO). The six subunits of the hexamer are shown in alternating slate gray and light gray. The di-ribonucleotides (5'-R(P*UP*C)-3') are shown in spacefill and colored CPK. ANPPNP is shown in spacefill yellow. Figure $24$ shows an interactive iCn3D model of the closed ring structure of the E. Coli Rho transcription termination factor in a complex with nucleic acid in the motor domains (2HT1). The six subunits of the hexamer are again shown in alternating slate gray and light gray. The ssRNAs are shown in spacefill with the backbones in one color and the bases in CPK colors. Figure $25$ shows an interactive iCn3D model of the E. coli Rho-dependent Transcription Pre-termination Complex (6XAS) (long load). • The RNA polymerase subunits (α2ββω) are shown in light cyan • The six subunits of the Rho hexamer are again shown in alternating slate gray and light gray • NusA is shown in magenta • Both DNA strands are shown in spacefill with the backbone blue and the bases red • Only part of the RNA is shown (spacefill, backbone yellow, bases CPK), so it appears discontinuous Eukaryotic Transcriptional Termination In eukaryotes, termination of protein-coding gene transcription by RNA polymerase II (Pol II) usually requires a functional polyadenylation (pA) signal, typically a variation of the AAUAAA hexamer. Nascent pre-mRNA is cleaved and the 5′ fragment is polyadenylated at the pA site shortly downstream from the hexamer by cleavage and pA factors (CPFs). Two mechanisms have been suggested for pA-dependent transcription termination. In the allosteric model, the pA signal and/or other termination signals bind with the pA signal downstream region (PDR) and induce reorganization of the Pol II complex. This includes the association or dissociation of endonuclease components such as the CPFs. This causes conformational changes in Pol II and TEC disassembly ensues. In the kinetic model, also known as the “torpedo” model, cleavage at the pA site separates the pre-mRNA from the TEC, which continues synthesizing a downstream nascent transcript. This new transcript is a substrate of XRN2/Rat1p, a processive 5′-to-3′ exoribonuclease that catches up with, and disassembles, the TEC by an unknown mechanism. The two pA-dependent models are not mutually exclusive, and unified models have been proposed. Loosely conserved pA signal sequences downstream of protein-coding genes bind to components of the polyadenylation factor (CF1) complex leading to the assembly of the cleavage and polyadenylation machinery. Termination is coupled to cleavage in a manner that has not yet been completely resolved, however, one of the major factors involved in yeast pA termination is the endonuclease, Ysh1. For example, the depletion of Ysh1 blocks TEC dissociation, but does not cause substantial readthrough at the termination site (Fig. 26.1.18 A&B). These results suggest that Ysh1 does not directly cause the pausing that occurs in the allosteric termination pathway, but rather plays a role in the dissociation of the Pol II complex from the DNA template, as shown in Figure $26$. It should be noted that not all pA-dependent termination is dependent on Ysh1 and that other mechanisms of pA-mediated termination still remain to be elucidated. Panel (A) shows the elongating Pol II (green) terminates pA transcripts (A) after an allosteric change (red) that reduces processivity. Panel (B) shows the depletion of Ysh1 leads to minimally extended readthrough transcripts but does not block the allosteric change in Pol II. Panel (C) shows how Nrd1 and Nab3 binding recruits Sen1 for termination of non-pA transcripts. Panel (D) shows Pol II elongation complex lacking Nrd1 does not recognize termination sequences in the nascent transcript and thus does not facilitate the allosteric transition in Pol II. This leads to a processive readthrough. Panel(E) shows how Nrd1 and Nab3 recognize terminator sequences allowing the allosteric change in Pol II but depletion of Sen1 blocks the removal of Pol II from the template. Figure from: The mechanisms of termination of Pol II-mediated transcription differ for coding and non-coding transcripts. Coding transcripts and possibly some stable uncharacterized transcripts (SUTs) are nearly always processed at the 3′-end by the cleavage and polyadenylation (pA) machinery and are processed by the pA-dependent termination mechanisms described above. In contrast, ncRNAs are terminated and processed by an alternative pathway that, in yeast, requires the RNA-binding proteins Nrd1 and Nab3, as well as, the RNA helicase Sen1. Nrd1 and Nab3 recognize RNA sequence elements downstream of snoRNAs and CUTs and this leads to the association of a complex that contains the DNA/RNA helicase Sen1 and the nuclear exosome. The nuclear exosome is a complex of ribonucleases with 3' to 5' exonuclease and endonuclease activity. It functions to degrade unstable or incorrect RNA transcripts. Both Nrd1 and Sen1 depletion lead to readthrough transcription of ncRNAs, suggesting their importance in non-pA-dependent transcription termination (Fig 26.1.18 C & D). Furthermore, depletion of Nrd1 also causes the accumulation of longer readthrough ncRNAs, suggesting its role in trafficking ncRNAs to the nuclear exosome following termination. 1. Parker, N., Schneegurt, M., Thi Tu, A-H., Lister, P., Forster, B.M. (2019) Microbiology. Openstax. Available at: https://opentextbc.ca/microbiologyopenstax/ 2. Palazzo, A., and Lee, E.S. (2015) Non-coding RNA: what is function and what is junk? Frontiers in Genetics 6:2 Available at: file:///C:/Users/flatt/AppData/Local/Temp/fgene-06-00002.pdf 3. Wikipedia contributors. (2020, July 9). RNA. In Wikipedia, The Free Encyclopedia. Retrieved 15:30, August 6, 2020, from https://en.Wikipedia.org/w/index.php?title=RNA&oldid=966784317 4. Burenina, O.Y., Oretskaya, T.S., and Kubareva, E.A. (2017) Non-Coding RNAs As Transcriptional Regulators in Eukaryotes. Acta Naturae 9(4):13-25. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5762824/ 5. Khatter, H., Vorlander, M.K., and Muller C.W. (2017) RNA polymerase I and III: similar yet unique. Current Opinion in Structural Biology 47:88-94. Available at: https://www.sciencedirect.com/science/article/pii/S0959440X17300313 6. Wikipedia contributors. (2020, May 8). Sigma factor. In Wikipedia, The Free Encyclopedia. Retrieved 17:50, August 7, 2020, from https://en.Wikipedia.org/w/index.php?title=Sigma_factor&oldid=955570499 7. Bae, B., Felkistov, A., Lass-Napiokowska, A., Landick, R., and Darst, S.A. (2015) Structure of a bacterial RNA polymerase holoenzyme open protomer complex. eLife 4:e08504. Available at: https://elifesciences.org/articles/08504 8. Petrenko, N., Jin, Y., Dong, L., Wong, K.H., and Struhl, K. (2019) Requirements for RNA polymerase II preinitiation complex formation in vivo. eLife 8:e43654. Available at: https://elifesciences.org/articles/43654 9. Gupta, K., Sari-Ak, D., Haffke, M., Trowitzsch, S., and Berger, I. (2016) Zooming in on transcription preinitiation. J Mol Biol. 428(12):2581-2591. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4906157/ 10. Wikipedia contributors. (2020, April 17). TATA-binding protein. In Wikipedia, The Free Encyclopedia. Retrieved 14:54, August 8, 2020, from https://en.Wikipedia.org/w/index.php?title=TATA-binding_protein&oldid=951583592 11. Patel, A.B., Greber, B.J., and Nogales, E. (2020) Recent insights into the structure of TFIID, its assembly, and its binding to core promoter. Curr Op Struct Bio 61:17-24. Available at: https://www.sciencedirect.com/science/article/pii/S0959440X19301113#fig0010 12. Ruff, E.F., Record, Jr., M.T., Artsimovitch, I., (2015) Initial events in bacterial transcription initiation. Biomolecules 5(2):1035-1062. Available at: https://www.mdpi.com/2218-273X/5/2/1035/htm 13. Kireeva, M., Opron, K., Seibold, S., Domecq, C., Cukier, R.I., Coulombe, B., Kashlev, M., and Burton, Z. (2102) Molecular dynamics and mutational analysis of the catalytic and translocation cycle of RNA polymerase. BMC Biophysics 5(1):11. Available at: https://www.researchgate.net/publication/225281979_Molecular_dynamics_and_mutational_analysis_of_the_catalytic_and_translocation_cycle_of_RNA_polymerase/figures?lo=1 14. Washburn, R.S., and Gottesman, M.E. (2015) Regulation of transcription elongation and termination. Biomolecules 5(2):1063-1078. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4496710/pdf/biomolecules-05-01063.pdf 15. Zenkin, N., and Yuzenkova, Y. (2015) New insights into the functions of transcription factors that bind the RNA polymerase secondary channel. Biomolecules 5(3):1195-1209. Available at: https://www.mdpi.com/2218-273X/5/3/1195/htm 16. Gocheva, V., LeGall, A., Boudvillain, M., Margeat, E., and Nollmann, M. (2015) Direct observation of the translocation mechanism of transcription termination factor Rho. Nuc Acids Res 43(1):10.1093. Available at: https://www.researchgate.net/publication/272162172_Direct_observation_of_the_translocation_mechanism_of_transcription_termination_factor_Rho 17. Miki, T.S., Carl, S.H., and Groβhans, H. (2017) Two disctinct transcription termination modes dictated by promoters. Genes & Dev 31:1-10. Available at: https://www.researchgate.net/publication/320350041_Two_distinct_transcription_termination_modes_dictated_by_promoters 18. Gurumurthy, A., Shen, Y., Gunn, E.M., Bungert, J. (2018) Phase separation and transcription regulation: Are Super-Enhancers and Locus Control Regions primary sites of transcription complex assembly? BioEssays 1800164. Available at: https://www.researchgate.net/publication/329331157_Phase_Separation_and_Transcription_Regulation_Are_Super-Enhancers_and_Locus_Control_Regions_Primary_Sites_of_Transcription_Complex_Assembly 19. Suñé-Pou, M., Prieto-Sánchez, Boyero-Corral, S., Moreno-Castro, C., El Yousfi, Y., Suñé-Negre, J.M., Hernández-Munain, C., and Suñé, C. (2017) Targeting splicing in the treatment of human disease. Genes 8(3):87. Available at: https://www.mdpi.com/2073-4425/8/3/87/htm 20. Schaughency, P., Merran, J., and Corden J.L. (2014) Genome-wide mapping of yeast RNA polymerase II termination. PLOS Genetics 10(10):e1004632 Available at: https://journals.plos.org/plosgenetics/article?id=10.1371/journal.pgen.1004632 21. Nourse, J., Spada, S., and Danckwardt, S. (2020) Emerging roles of RNA 3'-end cleavage and polyadenylation in pathogenesis, diagnosis, and therapy of human disorders. Biomolecules 10(6):915. Available at: https://www.mdpi.com/2218-273X/10/6/915/htm 22. Wikipedia contributors. (2020, July 30). Five-prime cap. In Wikipedia, The Free Encyclopedia. Retrieved 05:53, August 11, 2020, from https://en.Wikipedia.org/w/index.php?title=Five-prime_cap&oldid=970240533 23. Cortes, T. and Cox, R.A. (2015) Transcription and translation of the rpsJ, rplN and rRNA operons of the tubercle bacillus. Microbiology (2015) 161:719-728. Available at: https://www.microbiologyresearch.org/docserver/fulltext/micro/161/4/719_mic000037.pdf?expires=1597159574&id=id&accname=guest&checksum=6FFC9C066EF41C7799FAE843CE94C49F 24. Hein, P.P. and Landick, R. (2010) The bridge helix coordinates movements of modules in RNA polymerase. BMC Biology 8:141. Available at: https://bmcbiol.biomedcentral.com/articles/10.1186/1741-7007-8-141 25. Gonatopoulos-Pournatzis, T., and Cowling, V.H. (2014) Cap-binding complex (CBC). Biochem. J. 457:231-242. Available at: https://www.researchgate.net/publication/259392894_Cap-binding_complex_CBC

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/25%3A_RNA_Metabolism/25.01%3A_DNA-Dependent_Synthesis_of_RNA.txt

Search Fundamentals of Biochemistry Post-transcriptional modifications of rRNA and tRNA will be topics of Chapter 27 as their structure and function in protein synthesis will be a focal point. Thus, this section will focus on post-transcriptional modifications of mRNA. We'll spend most of our time on eukaryotic RNA processing. Prokaryotic RNA Processing First, let's take a brief look at a fascinating feature of transcription in bacterial cells. Bacterial cells do not have extensive post-transcriptional modifications of mRNA primarily because transcription and translation are coupled processes. Bacterial cells lack the physical barrier of a nucleus, which allows transcription and translation machinery to function at the same time, enabling the concurrent translation of an mRNA while it is being transcribed (Fig 26.2.1). As mRNA is synthesized in prokaryotes, a ribosome binding motif called the Shine-Dalgarno sequence, located in the 5' untranslated region of the mRNA emerges early, allowing the ribosome to bind and translation to occur. In addition, the protein N-utilzation Substance, better known as NusG, plays a critical role. NusG has three separate domains and the functions of two of them are known. The NusG N-terminal domain (NusG-NTD) can bind to RNAP, whereas the C-terminal domain (NusG-CTD) can combine with the NusE (RpsJ) component of ribosomes. These two functions of NusG enable transcription to be coupled with translation. NusG CTD can also bind to Rho to terminate transcription, as shown in Figure \(1\). Figure \(1\): The roles of NusG in transcription/translation coupling. (a) Composition of an active RNAP complex. RNAP is shown in dark grey, DNA in blue and nascent RNA in red. The ribosome is shown in green with the nascent polypeptide chain in light grey; the bulge in the small subunit denotes the location of NusE (RpsJ). NusG is shown in orange: its shape denotes two functional sections. The larger section denotes the N-terminal domain, which binds to RNAP. The smaller section denotes the C-terminal domain, which interacts with NusE in situ. Rho is shown in purple. (b) After the translation is completed, NusG remains bound to RNAP and may also bind to Rho through the C-terminal domain leading to the termination of transcription. Figure from: Cortes, T., and Cox, R.A. (2015) Microbiology 161:719-728. Another protein, NusA, slows RNAP, in contrast to NusG which increases its processivity, as reflected by the length of the RNA made before RNAP falls off. Eukaryotic RNA Processing In multicellular organisms, almost every cell contains the same genome, yet complex spatial and temporal diversity is observed in gene transcripts. This is achieved through multiple levels of processing leading from gene to protein, of which RNA processing is an essential stage. Following the transcription of a gene by RNA polymerases to produce a primary mRNA transcript, further processing is required to produce a stable and functional mature RNA product. This involves various processing steps including RNA cleavage at specific sites, intron removal, called splicing, which substantially increases the transcript repertoire, and the addition of a 5'CAP. Another crucial feature of the RNA processing of most genes is the generation of 3′ ends through an initial endonucleolytic cleavage, followed in most cases by the addition of a poly(A) tail, a process termed 3′ end cleavage and polyadenylation (CPA). Cleavage and 3'-Polyadenylation (CPA) Polyadenylation is a required step for the correct termination of nearly all mRNA transcripts. Except for replication-dependent histone genes, metazoan protein-encoding mRNAs contain a uniform 3' end consisting of a stretch of adenosines. In addition to determining the correct transcript length at transcription termination, the poly(A) tail helps to ensure the translocation of the nascent RNA molecule from the nucleus to the cytoplasm, enhances translation efficiency, acts as a signal feature for RNA degradation, and thereby contributes to the production efficiency of a protein. Cleavage and polyadenylation (CPA) are carried out by the cleavage/polyadenylation apparatus (CPA), a multi-subunit 3′ end processing complex, which involves over 80 proteins, comprised of four core protein subcomplexes, as shown in Figure \(2\). These consist of 1. cleavage and polyadenylation specificity factor (CPSF), comprised of proteins CPSF1-4, factor interacting with PAPOLA and CPSF1 (FIP1L1), and WD repeat domain 33 (WDR33) (shown in green below); 2. cleavage stimulation factor (CstF), a trimer of CSTF1-3 (shown in red below; 3. cleavage factor I (CFI), a tetramer of two small nudix hydrolase 21 (NUDT21) subunits, and two large subunits of CPSF7 and/or CPSF6 (shown in orange in Figure 26.2.2 A); Note: Nudix are named for nucleoside diphosphates hydroxylases. 4. cleavage factor II (CFII), composed of cleavage factor polyribonucleotide kinase subunit 1 (CLP1) and PCF11 cleavage and polyadenylation factor subunit (PCF11) (shown in yellow in Figure 26.2.2 A). Additional factors include symplekin, the poly(A) polymerase (PAP), and the nuclear poly(A) binding proteins such as poly(A) binding protein nuclear 1 (PABPN1). CPA is initiated by this complex recognizing specific sequences within the nascent pre-mRNA transcripts termed polyadenylation signals (PAS). The PAS sequence normally consists of either a canonical 6 base sequence, the AATAAA hexamer, or a close variant usually differing by a single nucleotide (e.g., ATTAAA, TATAAA). It is located 10 to 35 nucleotides upstream of the cleavage site (CS) usually consisting of a CA dinucleotide. The PAS is also determined by surrounding auxiliary elements, such as upstream U-rich elements (USE), or downstream U-rich and GU-rich elements and G-rich sequences (DSE). As soon as the nascent RNA molecule emerges from RNA polymerase II (RNA Pol II), the CPSF complex is recruited to the PAS AATAAA hexamer, through numerous interactions. Upon successful assembly of this macromolecular machinery, CPSF3 performs the endonucleolytic cleavage followed by a non-templated addition of approximately 50-100 A residues. Panel (A) shows the core 3′ end processing machinery consists of complexes composed of multiple trans-acting proteins interacting with RNA via multiple cis-elements (USE = upstream sequence element; PAS = poly(A) signal; CS = cleavage site; DSE = downstream sequence element; CTD = C-terminal domain). Upon co-transcriptional assembly of these complexes, RNA cleavage and polyadenylation occur to form the 3′ end of the nascent RNA molecule. Panel (B) shows more than 70% of all genes harbor more than one polyadenylation signal (PAS). This gives rise to transcript isoforms differing at the mRNA 3′ end. While alternative polyadenylation (APA) in 3′UTR changes the properties of the mRNA (stability, localization, translation), internal PAS usage (in introns or the coding sequence (CDS)) changes the C-termini of the encoded protein, resulting in different functional or regulatory properties. Alternative polyadenylation (APA) occurs when more than one PAS is present within a pre-mRNA and provides an additional level of complexity in CPA-mediated RNA processing (Figure 26.2.2 B). Early studies revealed a significant portion of genes undergo APA, and with the advent of next-generation RNA sequencing technologies, the large-scale regulation of genes has become apparent, with approximately 70% of the transcriptome exhibiting APA regulation. As APA determines 3′UTR content and thus the regulatory features available to the mRNA, changes in the APA profile of a gene can have enormous impacts on expression. For those trying to understand the structure and mechanism of the cleavage/polyadenylation apparatus (CPA), it is especially frustrating that different names are given to the constituents that comprise it, especially when comparing the proteins from different organisms. Hence it is useful to see multiple representations of the complex. Figure \(3\) shows a different cartoon representation of the cleavage and polyadenylation reactions. Note again the number of colored subcomplexes within the CPA as well as the different abbreviations shown for the individual proteins. This cartoon diagram is useful in visualizing the different steps involved. Figure \(3\): Cis-regulatory sequence elements and protein factors involved in cleavage and polyadenylation. Marsollier, A.-C.; Joubert, R.; Mariot, V.; Dumonceaux, J. Targeting the Polyadenylation Signal of Pre-mRNA: A New Gene Silencing Approach for Facioscapulohumeral Dystrophy. Int. J. Mol. Sci. 2018, 19, 1347. https://doi.org/10.3390/ijms19051347. Creative Commons Attribution License Panel (A) shows that the specificity and efficiency of 3′end processing are determined by the binding of more than 80 RNA-binding proteins to regulatory cis-acting RNA sequence elements including the polyadenylation signal (PAS) A[A/U]UAAA; the cleavage site (represented by NN) and the downstream sequence element (DSE). Auxiliary sequences can be found near the polyadenylation signal or the DSE. The core processing complex, which is sufficient for the cleavage and polyadenylation, is composed of approximately 20 proteins, distributed in 8 complexes: the cleavage and polyadenylation specificity factor (CPSF), the cleavage stimulation factor (CstF); the mammalian cleavage factors I (CFIm) and the mammalian cleavage factors II (CFIIm); the single protein poly(A) polymerase (PAP); the single protein poly(A)-binding protein nuclear 1 (PABPN1); the single protein RNA polymerase II large subunit (Pol II); and the symplekin. Subunits of the different factors are indicated. Panel (B) shows how CPSF and CstF are co-transcriptionally recruited to the poly(A) signal and the DSE respectively, causing an endonucleolytic cleavage of the pre-mRNA between the PAS and the DSE at the cleavage site. Two fragments are generated: one fragment with a free 5′phosphate group which is rapidly degraded by exoribonucleases and one fragment with a free 3′hydroxyl group on which 250 adenines will be added by PAP. The newly-synthetized poly(A) tail is covered by PAPBN1, allowing mRNA circularization and stabilization. Now let's look at the structure of some of the complexes of the cleavage/polyadenylation apparatus (CPA). In yeast, the 3' processing is carried out by the cleavage and polyadenylation factor (CPF) which is called the CPSF in humans. On endonuclease cleavage, the RNA bound to RNA polymerase II is in two pieces, as shown in the figure above. The main mRNA now has a 3'-OH which is the site of polyadenylation. The minor cleavage fragment has a 5'-phosphate which gets degraded by the exonuclease Rat1, which as it cleaves the minor product helps displaces RNA polymerase II and helps to stop transcription. In yeast, the CPF has 14 subunits with polymerase, nuclease, and phosphatase subcomplexes or "modules". The polymerase module, as the name implies, has the poly(A) polymerase, Pap1. Table \(1\) below shows some components of the polymerase module of both yeast CPF and human CPSF. yeast Polymerase Module of CPF Human mammalian polyadenylation specificity factor (mPSF) or CPSF Cft1 CPSF160 Pfs2 WDR33 Yth1 - RNA binding subunit CPSF30 - RNA binding subunit Fip1 - Pap1 binding subunit FIP1 - Pap1 binding subunit Table \(1\) Some components of the polymerase module of both yeast CPF and human CPSF. The nuclease module has an endonuclease (Ysh1) and a Mpe1 protein, which facilitate the cleavage site selection and polyadenylation. Table \(2\) below shows some components of the polymerase module in yeast and humans yeast human endonuclease Ysh1 endonuclease CPSF73 pseudo-nuclease Cft2 pseudo-nuclease CPSF100 multidomain protein Mpe1 multidomain protein RBBP6 Table \(2\): Some components of the polymerase module in yeast and humans Part of the Cft2 called the yeast polymerase module interacting motif (yPIM), as its name implies, interacts with the polymerase module, in part through the interaction of key and conserved aromatic residues in it (F537, Y549, and F558) with a hydrophobic binding site in Cft1 and Pfs1. These interactions are key in activating and regulating the endonuclease and polyadenylation activities and hence controlling the termination of transcription. Figure \(4\) shows an interactive iCn3D model of the yeast cleavage and polyadenylation specificity factor (CPF) polymerase module in complex with Mpe1, the yPIM of Cft2 and the pre-cleaved CYC1 RNA (7ZGR) The different parts of the complex are colored as shown below. • Yth1 (mRNA 3' processing protein): magenta • CFT1: green • MPE1: orange • Polyadenylation subunit 2 (Pfs2): yellow • cleavage factor 2 protein (CF2P): cyan • precleaved RNA CYC1: gray spacefill (backbone) with CPK-colored bases 5'-CAP Formation In eukaryotes, the 5′ cap, found on the 5′ end of the eventual mRNA molecule, consists of a guanine nucleotide connected to the mRNA via an unusual 5′ to 5′ triphosphate linkage, as shown in Figure \(5\). This guanosine is methylated on the 7 position directly after capping in vivo by a methyltransferase. It is referred to as the 7-methylguanylate cap, abbreviated m7G. In multicellular eukaryotes and some viruses, further modifications exist, including the methylation of the 2′ hydroxy-groups of the first 2 ribose sugars of the 5′ end of the mRNA. Cap-1 has a methylated 2′-hydroxy group on the first ribose sugar, while cap-2 has methylated 2′-hydroxy groups on the first two ribose sugars. The 5′ cap is chemically similar to the 3′ end of an RNA molecule (the 5′ carbon of the cap ribose is bonded, and the 3′-OH unbonded). This provides significant resistance to 5′ exonucleases. Small nuclear RNAs (snRNAs) contain unique 5′-caps. Sm-class snRNAs are found with 5′-trimethylguanosine caps, while Lsm-class snRNAs are found with 5′-monomethylphosphate caps. In bacteria, and potentially also in higher organisms, some RNAs are capped with NAD+, NADH, or 3′-dephospho-coenzyme A. In all organisms, mRNA molecules can be decapped in a process known as messenger RNA decapping. For capping with 7-methylguanylate, the capping enzyme complex (CEC) binds to RNA polymerase II before transcription starts. As soon as the 5′ end of the new transcript emerges from RNA polymerase II, the CEC carries out the capping process (this kind of mechanism ensures capping, as with polyadenylation). The enzymes for capping can only bind to RNA polymerase II that is engaging in mRNA transcription, ensuring the specificity of the m7G cap almost entirely to mRNA. The 5′ cap has four main functions: 1. Regulation of nuclear export 2. Prevention of degradation by exonucleases 3. Promotion of translation (see ribosome and translation) 4. Promotion of 5′ proximal intron excision In addition to the polyA tail, the nuclear export of RNA is regulated by the cap-binding complex (CBC), which binds to 7-methylguanylate-capped RNA, as shown in Figure \(6\). The CBC is then recognized by the nuclear pore complex and the mRNA is exported. Once in the cytoplasm after the pioneer round of translation, the CBC is replaced by the translation factors eIF4E and eIF4G of the eIF4F complex. This complex is then recognized by other translation initiation machinery including the ribosome, aiding in translation efficiency. Panel (a) CBC is required for pre-mRNA processing. The co-transcriptional binding of CBC to 7mG prevents the decapping activities of pre-mRNA degradation complexes [DXO (decapping exoribonuclease) and Dcp (decapping mRNA) Xrn2 (5′–3′ exoribonuclease 2)] and promotes pre-mRNA processing. CBC recruits P-TEFb [Cdk9/Cyclin T1 (CycT1)] to transcription initiation sites of specific genes promoting phosphorylation of the RNA pol II CTD at Ser2 residues. This results in the recruitment of splicing factors including SRSF1, which regulates both constitutive and alternative splicing events. Furthermore, CBC interacts with splicing machinery components that result in the spliceosomal assembly. CBC interacts with NELF and promotes pre-mRNA processing of replication-dependent histone transcripts. Panel (b) CBC forms a complex with Ars2 and promotes miRNA biogenesis by mediating pri-miRNA processing. Panel (c) CBC/Ars2 promotes pre-mRNA processing of replication-dependent histone transcripts. Panel (d) CBC promotes the export of U snRNA. CBC interacts with PHAX, which recruits export factors including CRM1 and RAN·GTP. Panel (e) CBC promotes the export of mRNA. For the export of transcripts over 300 nucleotides, hnRNP C interacts with CBC and inhibits the interaction between CBC and PHAX, allowing the CBC to interact with TREX and the transcript to be translocated to the cytoplasm. CBC interacts with the PARN deadenylase and inhibits its activity, protecting mRNAs from degradation. Panel (f) CBC mediates the pioneer round of translation. Cbp80 interacts with CTIF, which recruits the 40S ribosomal subunit via eIF3 to the 5′ end of the mRNA for translation initiation. Upon binding of importin-β (Imp-β) to importin-α (Imp-α), mRNA is released from CBC and binds to eIF4E for the initiation of the standard mode of translation. CBC-bound mRNP components not found in eIF4E-bound mRNPs are CTIF, exon junction complex (EJC), and PABPN1. Panel (g) The standard mode of translation is mediated by eIF4E cap-binding protein. eIF4E is a component of the eIF4F complex which promotes translation initiation. Capping with 7-methylguanylate prevents 5′ degradation in two ways. First, degradation of the mRNA by 5′ exonucleases is prevented by functionally looking like a 3′ end. Second, the CBC and eIF4E/eIF4G block the access of decapping enzymes to the cap. This increases the half-life of the mRNA, essential in eukaryotes as the export and translation processes take significant time. The mechanism that promotes the 5′ proximal intron excision during splicing is not well understood, but the 7-methylguanylate cap appears to loop around and interact with the spliceosome, potentially playing a role in the splicing process. The decapping of a 7-methylguanylate-capped mRNA is catalyzed by the decapping complex made up of at least Dcp1 and Dcp2, which must compete with eIF4E to bind the cap. Thus the 7-methylguanylate cap is a marker of an actively translating mRNA and is used by cells to regulate mRNA half-lives in response to new stimuli. During the decay process, mRNAs may be sent to P-bodies. P-bodies are granular foci within the cytoplasm that contain high levels of exonuclease activity. Triphosphatase and guanylyltransferase In capping the new mRNA, three different enzymes act sequentially: • A phosphatase cleaves a terminal phosphate from the 5' end which has 3 phosphates at the start leaving 2 phosphates (the yeast triphosphatase is called Cet1); • a GMP is added to the remaining diphosphate on the 5'-end to form a triphosphate in a reverse direction as shown in Figure 5 above (in yeast the RNA guanylyltransferase is called Ceg1); • a methyl group is added to the N7 of the guanine base by a methylase also called a methyl transferase These three enzymes are localized to a part of RNA polymerase that is highly phosphorylated, positioning them at the correct location for Cap formation. In yeast, the Figure \(7\) shows an interactive iCn3D model of the Saccharomyces cerevisiae Cet1 (the triphosphatase)-Ceg1 (the guanylyltransferase) mRNA Capping Apparatus (3KYH) The two enzymes exist as a heterotetramer of two homodimers. Two beta (Cet1-triphosphatase) subunits are shown in gray and the two alpha (Ceg1-guanylyltransferase) subunits are shown in cyan. The active site residues are shown in CPK-colored sticks and labeled. The two enzymes interact with yeast RNAP II mostly through Ceg1. Specifically, the Ceg1 oligonucleotide domain interacts with a motif, WAQKW (247-251), on Cet1. A conformational change in a flexible linker after that motif allows capping to ensue. Methyl transferase The next step is the methyl transfer to the N7 of guanine in the 5' end of the cap. Figure \(8\) shows an interactive iCn3D model of the Structure of a bacterial mRNA cap (Guanine-n7) methyltransferase (1RI1) 7-Methyl-guanosine-5'-triphosphate-5'-guanosine (GTG) is shown in sticks. The N7 nitrogen of guanine is labeled. S-Adenoslyl-L-homocysteine, the leaving group after methylation by S-adenosyl-L-methionine (SAM), is shown in spacefill. Key amino acid side chains in the active site are labeled (in small letters). The structure is most consistent with an in-line methyl transfer from SAM to the attacking nucleophile, the N7 of guanine. Specificity in most capping methylases occurs through noncovalent interactions (mostly base stacking, with two amino acid aromatic groups (an example of π-π stacking), or one aromatic and one nonpolar side chain. In the bacterial example shown above, the interactions include hydrophobic and hydrogen bonding interactions using Y284, F24, P175, E225, H144, and Y145. Decapping "What goes up must come down!" If the mRNA is capped during synthesis, there must be an enzyme to decap it. Figure \(9\) shows an interactive iCn3D model of the yeast mRNA decapping enzyme Dcp1-Dcp2 complex in the ATP bound closed conformation (2QKM) The rendering is colored as shown below: • a chain in cyan • b chain in gray • nudix motif in magenta • ATP in spacefill Dcp2p, as with many other enzymes, has an open (inactive) and closed (active) conformation suggesting that a conformational change between the two states regulates decapping. The ATP binding site demarcates the active site. the Dcp1 protein probably functions to stabilize the closed state. mRNA Splicing Eukaryotic genes that encode polypeptides are composed of coding sequences called exons (ex-on signifies that they are expressed) and intervening sequences called introns (int-ron denotes their intervening role). Transcribed RNA sequences corresponding to introns do not encode regions of the functional polypeptide and are removed from the pre-mRNA during processing. All of the intron-encoded RNA sequences must be completely and precisely removed from a pre-mRNA before protein synthesis so that the exon-encoded RNA sequences are properly joined together to code for a functional polypeptide. If the process errs by even a single nucleotide, the sequences of the rejoined exons would be shifted, and the resulting polypeptide would be nonfunctional. The process of removing intron-encoded RNA sequences and reconnecting those encoded by exons is called RNA splicing. Intron-encoded RNA sequences are removed from the pre-RNA while it is still in the nucleus. Although they are not translated, introns appear to have various functions, including gene regulation and mRNA transport. On completion of these modifications, the mature transcript, the mRNA that encodes a polypeptide, is transported out of the nucleus, destined for the cytoplasm for translation. Introns can be spliced out differently, resulting in various exons being included or excluded from the final mRNA product. This process is known as alternative splicing. The advantage of alternative splicing is that different types of mRNA transcripts can be generated, all derived from the same DNA sequence. In recent years, it has been shown that some archaea also can splice their pre-mRNA. The splicing reaction is catalyzed by the spliceosome, a macromolecular complex formed by five small nuclear ribonucleoproteins (snRNPs), termed U1, U2, U4, U5, and U6, and approximately 200 proteins, as shown in Figure \(10\). Each of these snRNPs contains snRNAs that can interact with each other through intrastrand hydrogen bonding and hence which help localize the snRNPs to the complexes. The assembly of the spliceosome on pre-mRNA includes the binding of U1 snRNP, U2 snRNP, the pre-formed U4/U6-U5 triple snRNP, and the Prp19 complex. This assembly occurs through the recognition of several sequence elements on the pre-mRNA that define the exon/intron boundaries, which include the 5′ and 3′ splice sites (SS), the associated 3′ sequences for intron excision, the polypyrimidine (Py) tract, and the branch point sequence (BPS). The assembly of the spliceosome during the process is depicted in Figure \(10\). In the first step of the splicing process, the 5′ splice site (GU, 5′ SS) is bound by the U1 snRNP, and the splicing factors SF1/BBP and U2AF cooperatively recognize the branch point sequence (BPS), the polypyrimidine (Py) tract, and the 3′ splice site (AG, 3′ SS) to assemble complex E. The binding of the U2 snRNP to the BPS results in the pre-spliceosomal complex A. Subsequent steps lead to the binding of the U4/U5–U6 tri-snRNP and the formation of complex B. Complex C is assembled after rearrangements that detach the U1 and U4 snRNPs to generate complex B*. Complex C is responsible for the two transesterification reactions at the SS. Additional rearrangements result in the excision of the intron, which is removed as a lariat RNA, and ligation of the exons. The U2, U5, and U6 snRNPs are then released from the complex and recycled for subsequent rounds of splicing There are different ways that pre-spliceosomes convert to full spliceosomes. The interactions of the small ribonucleoproteins U1 and U2, which bind at the 5' and 3' end of the introns respectively, are key. When multiple introns exist, a few different pathways occur. The interactions can be upstream (cross-introns) and downstream (cross-exons), as shown in Figure \(11\). Figure \(11\): Implications of synergistic U2 recruitment for the mechanism of exon and intron recognition. Braun et al. (2018) . Synergistic assembly of human pre-spliceosomes across introns and exons. eLife 7:e37751. https://doi.org/10.7554/eLife.37751. Creative Commons Attribution License The left-hand side of the figure shows how the interaction of U1 and U2 can occur across a single intron, to form the mature version of the spliceosome showing the U1-U2 interaction. The colored arrow shows possible interactions that either speed up or slow down the U1-U2 mature interactions. The right-hand side shows the formation of an intermediate cross-exon U2-U1 pair in which the exons that flank two introns are first delineated, which then leads to the physical U1-U2 interaction in the mature spliceosome when cross-intron interactions occur. Now let's move to show the actual structures of two of the spliceosome structures shown in Figure 10 above. Figure \(12\) shows an interactive iCn3D model of the human fully-assembled precatalytic spliceosome (pre-B complex) (6QX9) The full structure of the spliceosome is almost too complicated to understand even if displayed in a molecular model. It's better grasped in a way by the cartoon figure shown previously. To avoid unnecessary visual complexity, the protein subunits in the iCn3D model of the human spliceosome are shown in a gray cartoon, and only the RNA molecules in the Un small ribonucleoproteins are shown in color, as described below. The resolution is such that each nucleotide in the RNA polymers is shown just as a single-colored sphere not connected to the next nucleotide in the RNA. • U1 snRNA - magenta • U6 snRNA - orange • U5 snRNA - yellow • U2 snRNA - cyan • U4 snRNA - blue • AdML pre-mRNA - black Since it has U1, U2, U4, U5, and U6, it most closely resembles Complex B (or an immediate precursor pre-B of it) as shown in Figure 10. This form occurs before U1 snRNP dissociates. A helix from the 5'-single-stranded U1 snRNA inserts into a helicase in the complex between two RecA domains which bind ATP and unfold the nucleic acids. This allows the 5' single-stranded RNA to form interactions with an ACAGAGA sequence (mobile loop) in U6. These conformational changes allow the eventual separation of the U4 and U6 snRNA, freeing the snRNA in U6 is form the catalytic site. The B complex itself does not have a functioning active site. After the dissociation of U1 and U4 snRNPs, and the binding of another 20 or so proteins, the active spliceosome is formed. Now let's look at the final catalytic structure, Complex C, which contains only three of the snRibonucleoproteins, U2, U5, and U6. Figure \(13\) shows an interactive iCn3D model of the Human C Complex Spliceosome (7A5P) The color coding is as follows: • protein - gray • U6 snRNA - orange • U5 snRNA - yellow • U2 snRNA - cyan • AdML pre-mRNA - magenta The interactions of the snRNAs are key for structure and catalysis in the spliceosome, and in the figures above the interactions are had to discern. Figure \(14\) shows the interactions among U2, U5, and U6 snRNA in the human and yeast spliceosomes. Figure \(14\): The RNA elements and the splicing active site of the human mature Bact complex. Zhang, X., Yan, C., Zhan, X. et al. Structure of the human-activated spliceosome in three conformational states. Cell Res 28, 307–322 (2018). https://doi.org/10.1038/cr.2018.14. Creative Commons Attribution 4.0 Unported License. http://creativecommons.org/licenses/by/4.0/ Panel (A) shows the structure of the RNA elements in the core of the human mature Bact complex. The color-coded RNA elements of the human Bact complex are shown in the left panel, and their superposition with those of the S. cerevisiae Bact complex is displayed in the right panel. All yeast RNA elements are colored grey. The helix II of the U2/U6 duplex in the human Bact complex is bent relative to that in the yeast complex. Panel (B) shows a structural overlay of the active site RNA elements between the human and S. cerevisiae Bact complexes. Panel (C) shows that the U6/intron duplex in the human Bact complex is considerably longer than that in the S. cerevisiae Bact complex. In mammals, the first catalytic step of the splicing reaction begins when the U1 snRNP binds the 5′ SS of the intron (defined by the consensus sequence AGGURAGU), and the splicing factors SF1 and U2AF cooperatively recognize the BPS, Py, and 3′ SS to assembled complex E or the commitment complex. Subsequently, U2 snRNP and additional proteins are recruited to the pre-mRNA BPS to form the pre-spliceosome or complex A. The binding of the U4/U6-U5 tri-snRNP forms the pre-catalytic spliceosome or complex B. After RNA-RNA and RNA-protein rearrangements at the heart of the spliceosome, U1 and U4 are released to form the activated complex B or complex B* This complex is responsible for executing the first catalytic step, through which the phosphodiester bond at the 5′ SS of the intron is modified by the 2′-hydroxyl of adenosine of the BPS to form a free 5′ exon and a branched intron, as shown in Figure \(15\). The reaction of the 2'-hydroxyl from the branch point adenosine nucleotide is known as a transesterification reaction. During this process, additional rearrangements occur to generate the catalytic spliceosome or complex C (see Figure 10 above), which is responsible for catalyzing the second transesterification reaction leading to intron excision and exon–exon ligation. The resulting intron structure is referred to as a lariat structure. After the second catalytic step, the U2, U5, and U6 snRNPs are released from the post-spliceosomal complex and recycled for additional rounds of splicing. Panel (A) shows a schematic diagram of the pre-mRNA with exons and introns indicated. Key sequences are required for splicing at the 5' and 3' intron locations, and for the recognition and positioning of the branch point Adenosine residue for the first transesterification reaction. ( Panel (B) shows a schematic of the two transesterification reactions required for intron removal. The branch point 2'-OH residue mediates attack on the 5'-phosphate of the intron guanosine residue located at the 5'-splice site. This releases the 3' hydroxyl of Exon 1 which subsequently mediates the attack of the 5' phosphate of the first guanosine residue in Exon 2. The 3' hydroxyl of the intron guanine residue is released forming the Lariat structure and Exon 1 is ligated to Exon 2. Alternative Splicing (AS) offers an additional mechanism for regulating protein production and function. AS options are determined by the expression of or exposure to trans elements present within unique cellular locations and environments. Additional sequence elements within the mRNA, known as exonic and intronic splicing silencers or enhancers (ESS, ISS, ESE, and ISE, respectively), participate in the regulation of AS. Specific RNA-binding proteins, including heterogeneous nuclear ribonucleoproteins (hnRNPs) and serine/arginine-rich (SR) proteins, recognize these sequences to positively or negatively regulate AS, as shown in Figure \(16\). These regulators, together with an ever-increasing number of additional auxiliary factors, provide the basis for the specificity of this pre-mRNA processing event in different cellular locations within the body. The core cis sequence elements that define the exon/intron boundaries (5′ and 3′ splice sites (SS), GU-AG, polypyrimidine (Py) tract, and branch point sequence (BPS)) are poorly conserved. Additional enhancer and silencer elements in exons and introns (ESE: exonic splicing enhancers; ESI: exonic splicing silencers; ISE: intronic splicing enhancers; ISI: intronic splicing silencers) contribute to the specificity of AS regulation. Trans-acting splicing factors, such as SR family proteins and heterogeneous nuclear ribonucleoprotein particles (hnRNPs), bind to enhancers and silencers and interact with spliceosomal components. In general, SR proteins bound to enhancers facilitate exon definition, and hnRNPs inhibit this process. These trans-acting elements are expressed differentially within different locations or under different environmental stimuli to regulate AS. There are several different types of AS events, which can be classified into four main subgroups. The first type is exon skipping, which is the major AS event in higher eukaryotes. In this type of event, a cassette exon is removed from the pre-mRNA as shown in Figure \(17\) (panel A). The second and third types are alternative 3′ and 5′ SS selections (panels b and c). These types of AS events occur when the spliceosome recognizes two or more splice sites at one end of an exon. The fourth type is intron retention (panel d), in which an intron remains in the mature mRNA transcript. This AS event is much more common in plants, fungi, and protozoa than in vertebrates. Other events that affect the transcript isoform outcome include mutually exclusive exons (panel e), alternative promoter usage (panel f), and alternative polyadenylation (panel g). Here is a detailed and incredible narrated animation of splicing and the spliceosome.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/25%3A_RNA_Metabolism/25.02%3A_RNA_Processing.txt

Search Fundamentals of Biochemistry RNA Viruses Infections with RNA viruses place a constant burden on our healthcare systems and economy. Over the past century, this has been particularly true for infections with the Human immunodeficiency virus 1 (HIV-1), Influenza A virus (IAV), Rotavirus (RotaV), West Nile virus (WNV), Dengue virus (DV), Measles virus (MV), and the Porcine reproductive and respiratory syndrome virus (PRRSV). But also emergent RNA viruses can have considerable consequences, such as the Severe acute respiratory syndrome-related coronavirus (SARS-CoV) in 2003, the Middle East Respiratory Syndrome coronavirus (MERS-CoV), and most recently the Severe acute respiratory syndrome-related coronavirus-2 (SARS-CoV-2) in December of 2019. Eukaryotes and bacteria can be infected with a wide variety of RNA viruses. On average, these pathogens share little sequence similarity and use different replication and transcription strategies. RNA virus genomes can consist of double-stranded RNA (dsRNA) or single-stranded (ssRNA) as shown in Figure \(1\) (Panel a). In turn, the ssRNA viruses can be classified into positive sense (+RNA) and negative sense (−RNA) viruses, depending on the translatability of their genetic material. As summarized for four model RNA pathogens in Panel b, all RNA viruses use dedicated replication and transcription strategies to amplify their genetic material. The common denominator of these strategies is a conserved RNA-dependent polymerase domain. Figure \(1\) describes variations of the replication process in the different types of RNA viruses. Panel a shows a simplified taxonomy of the genome architecture of representative RNA viruses. Panel b (+RNA virus)shows infection with a +RNA virus—as exemplified here with a CoV-like virion—releases a single-stranded RNA genome into the cytoplasm (1). (2) Translation of the 5′-terminal open-reading frame of the genome produces the viral replicase. (3) This multi-enzyme complex includes RNA-dependent RNA polymerase (RdRp) activity (orange) and associates with intracellular membranes before −RNA synthesis commences. Newly synthesized −RNAs are subsequently used to produce new +RNAs (4), which are typically capped (yellow) and polyadenylated (polyA). (Retrovirus) HIV-1 genomes are packaged as ssRNA in virions. When the ssRNA is released (1) a cDNA copy is synthesized by the reverse transcriptase enzyme (RT) (2). The RNA is next degraded by the intrinsic RNase H activity in the RT (3) and the single-stranded cDNA is converted to dsDNA (4). The dsDNA is imported into the nucleus (5) for integration into the host’s genetic material. (−RNA virus) (1) As illustrated here with an IAV-like particle, infection with an −RNA virus releases a viral RNA genome that is associated with a viral polymerase (orange) and nucleoprotein (green). (2) In the case of non-segmented −RNA viruses, these complexes support transcription to produce viral mRNAs or cRNAs. (3) Viral mRNAs are next translated and new viral proteins complex with cRNAs to synthesize new vRNAs. (5) The vRNA-containing complexes of some segmented −RNA viruses are imported into the nucleus of the host cell, where (6) the RdRp produces mRNAs or cRNAs. (7) mRNAs are transported to the cytoplasm, while cRNAs are bound by new viral proteins to form cRNPs for −RNA synthesis. (dsRNA virus) Fully duplexed RNA genomes lack cap and polyA elements. (1) The RdRp (orange), therefore, transcribes the viral genome inside the capsid of the virion (blue and red), so viral mRNAs can be (2) released into the cytoplasm as illustrated here with a rotavirus-like virion. In the cytoplasm, the mRNA is translated (3) or replicated by newly synthesized viral RdRps. Viral RNA-dependent RNA Polymerases (RdRps) were discovered in the early 1960s from studies on mengovirus and polio viruses when it was observed that these viruses were not sensitive to actinomycin D, a drug that inhibits cellular DNA-directed RNA synthesis. This lack of sensitivity suggested that there is a virus-specific enzyme that could copy RNA from an RNA template and not from a DNA template. RdRps show some structural similarity to telomerase enzymes suggesting a potential ancestral relationship of telomerase with RdRps. The most famous example of RdRp is of the polio virus. The viral genome is composed of RNA, which enters the cell through receptor-mediated endocytosis. From there, the RNA can act as a template for complementary RNA synthesis, immediately. The complementary strand is then, itself, able to act as a template for the production of new viral genomes that are further packaged and released from the cell ready to infect more host cells. The advantage of this method of replication is that there is no DNA stage; replication is quick and easy. The disadvantage is that there is no 'backup' DNA copy (Fig 26.3.1b; + RNA virus). Reverse Transcriptase (RT) enzymes, on the other hand, that are common to retroviruses, convert the + RNA strand from the virus into a cDNA copy (Fig 26.3.1b; retroviruses). The RT enzyme then degrades the RNA and the single-stranded cDNA is converted to dsDNA. The dsDNA is then integrated into the host's genome where it can remain in a dormant state. Upon reactivation, + RNA will be manufactured, along with viral proteins used in the assembly of the infectious viral particles. Structure and Mechanism of RNA Viral Polymerases The RNA-dependent polymerase domain is a member of the superfamily of template-dependent nucleic acid polymerases and typically <400 amino acids in length. Its sequence is highly variable on average, with some regions showing less than ~10% conservation. Strong amino acid conservation can be observed, however, in regions that are directly involved in nucleotide selection or catalysis, such as the strictly conserved glycine and aspartates in the center of the domain. The prototypic RNA virus RNA polymerase domain harbors seven of such regions or motifs, which are arranged in the order G, F1–3, A, B, C, D, and E from amino- to carboxy-terminus. Each of the seven motifs in the RNA polymerase domain adopts a specific and conserved fold, as shown in Figure \(2\). Panel a shows the structure of the FMDV RdRp. The motifs A, B, C, D, E, F, and G are color-coded yellow, gold, orange, red, light green, aquamarine, and blue. Overall the polymerase structure adopts a shape that resembles a cupped right hand. Herein, motifs A–E lie on the palm, while motifs F and G are part of the fingers. In the side-view of the enzyme, the location of the template and NTP channels is indicated. Panel b shows conserved structural elements of the FMDV RdRp. Homomorphic A–G were mapped and color-coded yellow, gold, orange, red, light green, aquamarine, and blue, respectively. Images A and B are based on PDB accession 2E9R. Figure \(3\) shows an interactive iCn3D model of the Foot-and-mouth disease virus RNA-polymerase in complex with a template- primer RNA, PPi, and UTP (2E9Z) Active site region side chains are shown as sticks and labeled. The template stand is shown in yellow and the primer is in green. Note the conserved amino acids Tyr-336, the catalytic Asp-338, and Lys-387 in motifs C and E, respectively. The primer 3'OH forms an H bond with active site Asp 338 (on motif C). In some structures, Asp 338 is bound to a metal ion that interacts with the PPP of the incoming NTP. Two metal ions are involved in catalysis. Tyr 336 interacts with the primer nucleotide while Lys 387 and Arg 388 interact with the primer backbone. These three residues are highly conserved. The acceptor base of the template strands is adjacent to the NTP binding site. There is no proofreading activity in viral RNA replication. RNA-dependent RNA polymerases generally have a groove on top of the enzyme where RNA enters. It exits in the front. Nucleotides enter at a rear channel. This is illustrated in Figure \(4\) (Panel b). Following the convention for cellular DNA-dependent DNA polymerases (DdDp), the seven motifs and homomorphs are grouped into three subdomains. These subdomains are called fingers, palm, and thumb in reference to the polymerase domain’s likeliness to a cupped right hand, as shown in Figure \(4\) (Panel A). The finger subdomain loops that interconnect the fingers with the thumb in the RNA-dependent RNA polymerases (RdRps) of +RNA and dsRNA viruses create an overall “closed-hand” conformation that is unique to RNA-dependent RNA polymerases (RdRps) and generally not seen in crystal structures of DNA-dependent DNA polymerases (DdDps) or reverse transcriptases (RTs). The three subdomains of the RNA-dependent polymerases work together to facilitate the binding of RNA and nucleotides (NTPs). The thumb subdomain contains various residues that are involved in RNA binding and these generally pack into the minor groove of the template RNA. In some polymerases, the thumb also contains sequences that protrude into the template channel to help stabilize the initiating NTPs on the ssRNA template. Crucially, these protrusions are also able to undergo relatively large conformational rearrangements to facilitate the translocation of the template following the first condensation reaction. The other residues of the thumb subdomain contribute to the formation of the NTP tunnel, which sits at an ~110° angle with the template channel as shown in Figure \(4\), Panel A. The cavity is lined with positively charged amino acids, though charge interactions are likely not sufficient to guide NTPs into the interior. Indeed, molecular dynamics (MD) simulations have shown that the residues of the NTP channel can also explore a relatively large amount of space, which may allow the RdRp to actively “pump” NTPs down the channel. Panel (A) shows a ribbon representation of a typical picornaviral RdRP (model from the cardiovirus EMCV 3Dpol, PDB id. 4NZ0). The seven conserved motifs are indicated in different colors: motif A, red; motif B, green; motif C, yellow; motif D, sand; motif E, cyan; motif F, blue; motif G, pink; Panel (B) shows a lateral view of a surface representation of the enzyme (grey) that has been cut to expose the three channels that are the entry and exit sites of the different substrates and reaction products. The structural elements that support motifs A–G are also shown as ribbons. This panel also shows the organization of the palm sub-domain with motif A shown in two alternative conformations: the standard conformation (PDB id. 4NZ0) found in the apo-form of most crystallized 3Dpol proteins and the altered conformation found in the tetragonal crystal form of the EMCV enzyme (PDB id. 4NYZ). The alterations affect mainly Asp240, the amino acid in charge of incoming ribonucleotide triphosphate (rNTP) selection, and the neighboring Phe239 that move ~10 Å away from its position in the enzyme catalytic cavity directed towards the entrance of the nucleotide channel, approaching to motif F; Panel (C) shows a close-up of the structural superimposition of the two alternative conformations of the EMCV motif A; Panel (D) shows the PV replication-elongation complexes. Sequential structures illustrating the movement of the different palm residues from a binary PV 3Dpol-RNA open complex (left) to an open 3Dpol-RNA-rNTP ternary complex (middle) where the incoming rNTP is positioned in the active site for catalysis and, a closed ternary complex (right) after nucleotide incorporation and pyrophosphate (PPi) release. The residues DA (involved in rNTP selection through an interaction with the 2′ hydroxyl group), DC (the catalytic aspartate of motif C), KD (the general acid residue of motif D that can coordinate the export of the PPi group), and NB (a conserved Asn of motif B, interacting with DA) have been highlighted as sticks. The different structures correspond to the 3Dpol-RNA (PDB id. 3OL6), 3Dpol-RNA-CTP open complex (PDB id. 3OLB) and 3Dpol-RNA-CTP closed complex (PDB id. 3OL7) structures of PV elongation complexes, respectively The finger subdomain residues mainly pack into the major groove of the RNA template. Furthermore, upon entry of the template, they can unstack the single strand at position +3, as shown in Figure \(5\) (Panel A). The non-conserved structural elements of the fingers subdomain play a role in RNA binding as well. In particular, the fingertips of dsRNA and some +RNA virus RdRps allow the polymerase to “hold” the template without extensive conformational changes. The variations and extensions in the fingers subdomain has also been shown to play roles in protein–protein interactions, phosphorylation, oligomerization, and nuclear import. In contrast, the HIV-1 RT lacks such extensions and adopts a more “open-hand” or U-shaped binding cleft. As a consequence, the RT structure must rearrange its subdomains to coordinate the binding of the template, nascent strand, and incoming dNTPs. Panel a shows the structure of the PV active site as it moves from a native state or elongation complex (i) to an open complex (ii), and a closed complex (iii). The closed complex depicted here shows the active site after catalysis. Highlighted are the metal-binding aspartates of motifs A and C, and the lysine of motif D that acts as a general acid. Color coding by motif as in Fig. 26.3.2. Image based on PDB accessions 3Ol6, 3OLB, and 3OL7. Panel b shows a schematic presentation of the RdRp active site. The aspartates (Asp) of motif A (yellow) and C (orange) bind divalent metal ions (marked Mg and shaded grey), which are used to coordinate the formation of a new phosphodiester bond at the 3′-OH (red in panel ii) of the nascent strand (yellow). The general acid (red Lys/His in panel ii) is positioned near the β-phosphate of the incoming NTP to protonate the PPi leaving group. Panel c shows a simplified schematic of the kinetic steps of RNA polymerases. Asterisk indicates a closed complex The NTP and template entry channels meet at the palm subdomain - Figure \(5\), Panel A. This is a structure that is comprised of a central, partially formed three-stranded β-sheet, which is also present in RNA-recognition motifs (RRMs). The relative movement of these strands within the RRM is vital to catalysis and dependent on NTP binding. Only when a correct NTP binds can motif A and motif C align and the RRM fold be completed. This catalytically competent conformation of the active site is often referred to as the closed complex (not to be confused with the “closed-hand”, which refers to the overall structure of the RdRp) - Figure \(5\), Panel d. The polymerase reaction creates new phosphodiester bonds between NTPs using RNA as a template. The NTP substrates involved in this reaction are coordinated by two metal ions, which are bound by the conserved aspartates of motifs A and C - Figure \(5\) - panel a, i. They also position the NTP’s triphosphate optimally for attack by the sugar moiety of the nascent strand once its 3′ carbon has lost a proton- Figure \(5\), panel b. The N-terminal aspartate of motif C thus uses a metal ion to fix the α-phosphate of the incoming NTP and reduce the pKa of the nascent RNA’s 3′-OH to facilitate the attack - Figure \(5\), panel b, ii. The amino-terminal carboxylate of motif A, on the other hand, stabilizes the β- and γ-phosphates with its divalent metal as well as the pentavalent (phosphorane) intermediate that forms during catalysis - Figure \(5\), panel b, ii. Structural and biochemical analyses have shown that the formation of this transient structure is dependent on the attack of the NTP’s α-phosphate by the 3′-OH, which is often the rate-limiting catalytic step in NTP condensation - Figure \(5\), panel b. Motif D’s lysine or histidine assists the N-terminal aspartate of motif A in coordinating the β-phosphate of the incoming NTP, analogous to the trigger loop in DdDps. However, when the positively charged side chain of motif D approaches the β-phosphate, it can protonate the PPi leaving group as well - Figure \(5\), panel b, ii. This second protonation step in the active site is not essential for the polymerase reaction, since catalysis can still take place when motif D’s lysine has been mutated to a residue with a different pKa. The polymerase reaction rate will nevertheless be 50- to 1,000-fold higher when a lysine or histidine is present. Recent data even suggests that motif D can coordinate the export of the PPi group from the active site once catalysis has taken place, thereby triggering the translocation of the RNA. Overall, the RNA replication process can be summarized with this four-step mechanism: 1. Nucleoside triphosphate (NTP) binding - initially, the RdRp presents with a vacant active site in which an NTP binds, complementary to the corresponding nucleotide on the template strand. Correct NTP binding causes the RdRp to undergo a conformational change. 2. Active site closure - the conformational change, initiated by the correct NTP binding, results in the restriction of active site access and produces a catalytically competent state. 3. Phosphodiester bond formation - two Mg2+ ions are present in the catalytically active state and arrange themselves in such a way around the newly synthesized RNA chain that the substrate NTP can undergo a phosphatidyl transfer and form a phosphodiester bond with the newly synthesized chain. With the use of these Mg2+ ions, the active site is no longer catalytically stable, and the RdRp complex changes to an open conformation. 4. Translocation - once the active site is open, the RNA template strand can move by one position through the RdRp protein complex and continue chain elongation by binding a new NTP, unless otherwise specified by the template. Figure \(6\) shows a more detailed representation of the elongation catalytic cycle. Figure \(6\): Elongation catalytic cycle of RNA-dependent RNA polymerases RNA synthesis can be performed using a primer-independent (de novo) or a primer-dependent mechanism that utilizes a viral protein genome-linked (VPg) primer. The de novo initiation consists of the addition of a nucleoside triphosphate (NTP) to the 3'-OH of the first initiating NTP. During the following so-called elongation phase, this nucleotidyl transfer reaction is repeated with subsequent NTPs to generate the complementary RNA product. The termination of the nascent RNA chain produced by RdRp is not completely known, however, it has been shown that RdRp termination is sequence-independent. One feature of RNA-dependent RNA polymerase replication is the immense error rate during transcription. RdRps and RTs are known to have a lack of fidelity on the order of 104 nucleotides, which is thought to be a direct result of their insufficient proofreading abilities. This high rate of variation is favored in viral genomes as it allows for the pathogen to overcome defenses developed by hosts trying to avoid infection allowing for evolutionary growth. Let's look at another RdRp from the poliovirus. The virus has a single-stranded, positive-sense RNA genome, which makes it infectious in itself but much more infectious when replicated. It is translated into a polyprotein which is cleaved into about 12 separate proteins. One protein, 3Dpol, is an RNA-dependent RNA polymerase that transcribes the infecting +RNA strand into a -RNA strand, which then serves as a template for more +RNA strands. Figure \(7\) shows an interactive iCn3D model of the Poliovirus polymerase elongation complex with 2',3'-dideoxy-CTP (3OLB). The model shows the palm, finger, and thumb domains. Color coding is as follows: • red: finger domain • pink: pinky finger part of the finger domain • blue: thumb domain • light blue: primer grip at the beginning of the thumb domain • cyan: RNA template • green: RNA product • gray: palm domain Structural studies suggest that the pinky finger is involved in the initiation, while nucleotide binding and catalysis used the palm domain. The thumb domain appears to affect the translocation step. In most nucleases, the finger binds and positions NTPs in the active site. After catalysis, the reverse motion opens the active site which allows translocation by effectively ratcheting the template by one base pair which is driven by PPi release. These conformational changes can't be made so easily in +RNA stranded viral RdRps.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/25%3A_RNA_Metabolism/25.03%3A_RNA-Dependent_Synthesis_of_RNA_and_DNA.txt

<h210.6refs">26.4.6 References 1. Parker, N., Schneegurt, M., Thi Tu, A-H., Lister, P., Forster, B.M. (2019) Microbiology. Openstax. Available at: https://opentextbc.ca/microbiologyopenstax/ 2. Palazzo, A., and Lee, E.S. (2015) Non-coding RNA: what is function and what is junk? Frontiers in Genetics 6:2 Available at: file:///C:/Users/flatt/AppData/Local/Temp/fgene-06-00002.pdf 3. Wikipedia contributors. (2020, July 9). RNA. In Wikipedia, The Free Encyclopedia. Retrieved 15:30, August 6, 2020, from https://en.Wikipedia.org/w/index.php?title=RNA&oldid=966784317 4. Burenina, O.Y., Oretskaya, T.S., and Kubareva, E.A. (2017) Non-Coding RNAs As Transcriptional Regulators in Eukaryotes. Acta Naturae 9(4):13-25. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5762824/ 5. Khatter, H., Vorlander, M.K., and Muller C.W. (2017) RNA polymerase I and III: similar yet unique. Current Opinion in Structural Biology 47:88-94. Available at: https://www.sciencedirect.com/science/article/pii/S0959440X17300313 6. Wikipedia contributors. (2020, May 8). Sigma factor. In Wikipedia, The Free Encyclopedia. Retrieved 17:50, August 7, 2020, from https://en.Wikipedia.org/w/index.php?title=Sigma_factor&oldid=955570499 7. Bae, B., Felkistov, A., Lass-Napiokowska, A., Landick, R., and Darst, S.A. (2015) Structure of a bacterial RNA polymerase holoenzyme open protomer complex. eLife 4:e08504. Available at: https://elifesciences.org/articles/08504 8. Petrenko, N., Jin, Y., Dong, L., Wong, K.H., and Struhl, K. (2019) Requirements for RNA polymerase II preinitiation complex formation in vivo. eLife 8:e43654. Available at: https://elifesciences.org/articles/43654 9. Gupta, K., Sari-Ak, D., Haffke, M., Trowitzsch, S., and Berger, I. (2016) Zooming in on transcription preinitiation. J Mol Biol. 428(12):2581-2591. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4906157/ 10. Wikipedia contributors. (2020, April 17). TATA-binding protein. In Wikipedia, The Free Encyclopedia. Retrieved 14:54, August 8, 2020, from https://en.Wikipedia.org/w/index.php?title=TATA-binding_protein&oldid=951583592 11. Patel, A.B., Greber, B.J., and Nogales, E. (2020) Recent insights into the structure of TFIID, its assembly, and its binding to core promoter. Curr Op Struct Bio 61:17-24. Available at: https://www.sciencedirect.com/science/article/pii/S0959440X19301113#fig0010 12. Ruff, E.F., Record, Jr., M.T., Artsimovitch, I., (2015) Initial events in bacterial transcription initiation. Biomolecules 5(2):1035-1062. Available at: https://www.mdpi.com/2218-273X/5/2/1035/htm 13. Kireeva, M., Opron, K., Seibold, S., Domecq, C., Cukier, R.I., Coulombe, B., Kashlev, M., and Burton, Z. (2102) Molecular dynamics and mutational analysis of the catalytic and translocation cycle of RNA polymerase. BMC Biophysics 5(1):11. Available at: https://www.researchgate.net/publication/225281979_Molecular_dynamics_and_mutational_analysis_of_the_catalytic_and_translocation_cycle_of_RNA_polymerase/figures?lo=1 14. Washburn, R.S., and Gottesman, M.E. (2015) Regulation of transcription elongation and termination. Biomolecules 5(2):1063-1078. Available at: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4496710/pdf/biomolecules-05-01063.pdf 15. Zenkin, N., and Yuzenkova, Y. (2015) New insights into the functions of transcription factors that bind the RNA polymerase secondary channel. Biomolecules 5(3):1195-1209. Available at: https://www.mdpi.com/2218-273X/5/3/1195/htm 16. Gocheva, V., LeGall, A., Boudvillain, M., Margeat, E., and Nollmann, M. (2015) Direct observation of the translocation mechanism of transcription termination factor Rho. Nuc Acids Res 43(1):10.1093. Available at: https://www.researchgate.net/publication/272162172_Direct_observation_of_the_translocation_mechanism_of_transcription_termination_factor_Rho 17. Miki, T.S., Carl, S.H., and Groβhans, H. (2017) Two disctinct transcription termination modes dictated by promoters. Genes & Dev 31:1-10. Available at: https://www.researchgate.net/publication/320350041_Two_distinct_transcription_termination_modes_dictated_by_promoters 18. Gurumurthy, A., Shen, Y., Gunn, E.M., Bungert, J. (2018) Phase separation and transcription regulation: Are Super-Enhancers and Locus Control Regions primary sites of transcription complex assembly? BioEssays 1800164. Available at: https://www.researchgate.net/publication/329331157_Phase_Separation_and_Transcription_Regulation_Are_Super-Enhancers_and_Locus_Control_Regions_Primary_Sites_of_Transcription_Complex_Assembly 19. Suñé-Pou, M., Prieto-Sánchez, Boyero-Corral, S., Moreno-Castro, C., El Yousfi, Y., Suñé-Negre, J.M., Hernández-Munain, C., and Suñé, C. (2017) Targeting splicing in the treatment of human disease. Genes 8(3):87. Available at: https://www.mdpi.com/2073-4425/8/3/87/htm 20. Schaughency, P., Merran, J., and Corden J.L. (2014) Genome-wide mapping of yeast RNA polymerase II termination. PLOS Genetics 10(10):e1004632 Available at: https://journals.plos.org/plosgenetics/article?id=10.1371/journal.pgen.1004632 21. Nourse, J., Spada, S., and Danckwardt, S. (2020) Emerging roles of RNA 3'-end cleavage and polyadenylation in pathogenesis, diagnosis, and therapy of human disorders. Biomolecules 10(6):915. Available at: https://www.mdpi.com/2218-273X/10/6/915/htm 22. Wikipedia contributors. (2020, July 30). Five-prime cap. In Wikipedia, The Free Encyclopedia. Retrieved 05:53, August 11, 2020, from https://en.Wikipedia.org/w/index.php?title=Five-prime_cap&oldid=970240533 23. Cortes, T. and Cox, R.A. (2015) Transcription and translation of the rpsJ, rplN and rRNA operons of the tubercle bacillus. Microbiology (2015) 161:719-728. Available at: https://www.microbiologyresearch.org/docserver/fulltext/micro/161/4/719_mic000037.pdf?expires=1597159574&id=id&accname=guest&checksum=6FFC9C066EF41C7799FAE843CE94C49F 24. Hein, P.P. and Landick, R. (2010) The bridge helix coordinates movements of modules in RNA polymerase. BMC Biology 8:141. Available at: https://bmcbiol.biomedcentral.com/articles/10.1186/1741-7007-8-141 25. Gonatopoulos-Pournatzis, T., and Cowling, V.H. (2014) Cap-binding complex (CBC). Biochem. J. 457:231-242. Available at: https://www.researchgate.net/publication/259392894_Cap-binding_complex_CBC

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/25%3A_RNA_Metabolism/25.04%3A_26.4_References.txt

Search Fundamentals of Biochemistry Overview of Translation Within this chapter, we will cover the details of prokaryotic and eukaryotic translation. Translation is the process of converting the information housed in mRNA into the protein sequence. Essentially, you are translating the language of nucleotides into the language of amino acids. Recall that prokaryotic and eukaryotic transcription and translation systems differ in large part due to the compartmentalization of larger eukaryotic cells. Due to this compartmentalization, transcription and translation are separated spatially and temporally within the cell. Transcription occurs within the nucleus of eukaryotes and translation occurs within the cytoplasm. Prokaryotes do not have compartmentalization and have, thus, evolved a coupled transcription/translation system where both processes occur simultaneously. Both are illustrated in Figure \(1\). Panel (a) shows that prokaryotes lack cellular compartmentalization and show coupled transcription-translation processing; Panel (b) shows that eukaryotes have a high degree of compartmentalization and separate the processes of transcription, which is in the nucleus of the cell, from the processes of translation, which is localized in the cytoplasm. Recall that peptide formation is a dehydration reaction that combines the carboxylic acid of the upstream amino acid with the amine functional group of the downstream amino acid to form an amide linkage as shown in Figure \(2\). Water is the by-product. The ribosome (a large complex of peptides and rRNA molecules) serves as the enzyme that mediates this reaction. It requires a mature mRNA to serve as the template and directionally performs peptide bond synthesis from the N to the C-terminal of the growing peptide/protein. This is known as N- to C-synthesis. Note that the overall protonation state shown is very unlikely since under conditions when the carboxyl groups are protonated, so would the amines. This representation makes it easier to highlight the departing water To maintain proper protein function, the error rate of translation is approximately 10-4 or 1 error in every 10,000 amino acids encoded. The fidelity of protein synthesis is maintained by the ribosome's ability to match the code from the template mRNA stand with the appropriate amino acid. Template mRNA is read by the ribosome in groups of three nucleotides, called a codon, as shown in Figure \(3\). The template is non-overlapping and reads in discrete groups of three. This is known as the reading frame of the mRNA, and it is always read from the 5′ to 3′ direction. Thus, for each mRNA, there are three potential reading frames (panel B). Only one reading frame will be the correct one for protein synthesis. The ribosome must recognize and align the correct reading frame of the mRNA such that the correct codon sequences can be read. Small distinct tRNA molecules are tethered with specific amino acids and contain specific anticodons that complement mRNA codon sequences. The tRNA molecules can cycle on and off of the ribosome structure to hybridize with the correct codon sequences and chaperone the correct amino acid for peptide bond formation. The ribosome then serves as a ribozyme and mediates the peptidyl transferase activity to form the peptide bond. The mRNA is then shifted to reveal the next codon within the sequence and the process is repeated until the entire protein has formed. Panel C shows the codon chart for all of the possible combinations of three nucleotides. 64 possible codon combinations are possible using the 4 nucleotide possibilities, but only 20 amino acids are encoded during protein synthesis. Each codon is specific for a single amino acid. There is very little ambiguity within the code. However, there is redundancy within the code; i.e. many amino acids have more than one codon that encodes for that specific amino acid. To account for this redundancy, many tRNA molecules can recognize more than one codon using a single anticodon. This is known as degeneracy. Degeneracy usually occurs at the third position of the codon and is known as the wobble base position. Degeneracy helps to minimize the effects of mutations within the coding sequence, as mutations in the wobble base position will often lead to silent mutation– ie the mutation will still encode for the same amino acid. In addition, if comparing the polarity of amino acids encoded by the different codons, neighboring codons typically encode for amino acids with similar polarity, as shown in Figure \(4\). This also helps to minimize the effects of mutations, by converting one amino acid within the sequence to one that has similar polarity. This type of mutation is more likely to cause less disturbance to the 3-dimensional structure of the resulting protein and retain biological function. Degeneracy within the genetic code also allows for differential A/T & G/C concentrations within species. For example, the G/C content of bacteria can range from as low as 30% to as high as 70%. Organisms living at high temperatures or extreme environments often have higher G/C content. This effectively increases the hydrogen bond strength between the strands of the DNA (G/C pairs have 3 H-bonds, whereas A/T pairs only have 2) and causes an increase in the melting temperature of the chromosome. Thus, the DNA is stable at a higher temperature or under more extreme ionic conditions, such as high salt. A more detailed discussion of how a single tRNA can function to recognize more than one codon is the topic of the next section. The Genetic Code is universal for almost all species alive on the planet, providing support for a single origin of life. Most deviations in the code occur within the mitochondria of eukaryotic species, as shown in Figure \(5\). Transfer RNA (tRNA) Structure Transfer RNAs (tRNAs) are central players in translation, functioning as adapter molecules between the informational level of nucleic acids and the functional level of proteins. Typically, tRNA molecules are between 76 – 90 nucleotides long and show a highly conserved secondary and tertiary structure. They also show the highest amount of nucleotide modification of all types of RNA with modifications concentrated in two hotspots—the anticodon loop and the tRNA core region, where the D- and T-loop interact with each other, stabilizing the overall structure of the molecule, as shown in Figure \(6\). These modifications can cause large rearrangements as well as local fine-tuning in the 3D structure of a tRNA. The life of a transfer RNA (tRNA) molecule starts with a series of important maturation steps that can vary in their sequential order from case to case. Leader and trailer sequences are removed by a set of endo- and exonucleases, and in several tRNA precursors, splicing reactions excise intronic sequences. Furthermore, in many organisms, the sequence CCA, which represents the site of amino acid attachment, is not encoded but has to be added post-transcriptionally by CCA-adding enzymes. While all primary tRNA transcripts are composed of the four standard RNA bases A, C, G, and U, many of these nucleotides are modified, altering their properties in very different ways. Currently, 93 post-transcriptional modifications are known, and the variety of their functions is at least similarly diverse and not fully understood. The complexity of such modifications ranges from simple methylations at the bases or the ribose to rather complex and large base hypermodifications, whose synthesis often requires a whole cascade of enzymatic reactions. Modifications can alter a tRNA’s shape in subtle ways, but can also lead to massive structural rearrangements. In addition, they ensure efficient translation by maintaining the anticodon loop structure and promoting correct codon-anticodon interactions. After maturation, tRNAs have multiple interaction partners in their life cycle, ranging from aminoacyl-tRNA-synthetases that are responsible for amino acid attachment, to translation factors, ribosomes, and mRNAs. Apart from synthetases, these interaction partners do not specifically act on one individual tRNA transcript or isoacceptor, but on all tRNAs, similar to the above-mentioned CCA-adding enzyme. Thus, despite a high sequence variation, a cell’s tRNAs show a well-conserved cloverleaf-like secondary structure that was originally discovered in 1965. The similar structure of all tRNA molecules allows them to bind to common protein synthesis machinery, such as the ribosome and CCA-adding enzymes. The cloverleaf consists of five parts: the acceptor stem (containing the tRNA’s 5′- and 3′-ends), the D-arm, the anticodon arm, the variable loop, and the TΨC-arm (T-arm). At the 3′-terminus, the tRNA carries the CCA sequence, required for aminoacylation, tRNA positioning in the ribosome, and translation termination. In a conserved network of tertiary interactions, mostly between D- and T-loop, tRNAs fold into an L-shaped three-dimensional structure, which was first solved by Kim et al. in 1974, as shown in Figure \(6\) (Panel B). The anticodon and the amino acid-accepting CCA-ends are separated by the longest possible distance from each other. This conserved structure of a tRNA is essential for its recognition by the ribosome, other RNAs, and proteins and, consequently, for its functionality. For example, the CCA-adding enzyme uses the acceptor domain for substrate recognition, whereas aminoacyl-tRNA-synthetases use several recognition elements like anticodon, acceptor stem, or the discriminator position. Panel (A) shows the canonical cloverleaf secondary structure of cytosolic tRNAPhe from S. cerevisiae is shown with acceptor stem (blue), D-arm (green), anticodon arm (red), variable loop (purple) and TΨC-arm (yellow). The anticodon is labeled in grey, the discriminator base in orange and post-transcriptional modifications in red. Grey dashed lines indicate tertiary interactions based on structural data and the length of the RNA is indicated in parenthesis; Panel (B) shows the L-shaped tertiary structure of the cytosolic tRNAPhe from S. cerevisiae. Protein Data Bank entry (PDB): 1EHZ. The acceptor domain is composed of a stacked T-arm and acceptor stem, whereas D- and anticodon arm form the anticodon domain. The region where both domains come together and interact with each other via tertiary base pairing is also called the elbow region; Panel(C) shows the secondary structure of human mitochondrial tRNASer1, which lacks the whole D-arm; Panel (D) shows the secondary structure of the mitochondrial tRNAArg from the nematode Romanomermis culicivorax, which lacks both D- and T-arm. Instead, we find a so-called replacement loop. It represents the shortest tRNA found in vivo. Surprisingly, not all tRNAs fold into the canonical cloverleaf structure. Especially many mitochondrial tRNAs are reduced in length and sometimes completely lack the D- or T-arm as shown in Figure \(6\), Panel C. In the mitochondria of nematodes, this situation is carried to an extreme, as tRNAs lacking one or even both arms seem to be the rule (Panel D). Figure \(7\) shows an interactive iCn3D model of the yeast phenylalanine tRNA (1EHZ). The coloring, which matches those in Figure 6 above, is shown below: • Acceptor Stem - blue • D Arm - green • Anticodon= magenta • Variable Loop - pink • Tω C - yellow Post-transcriptional enzyme-catalyzed modification of tRNA occurs at many base and sugar positions and influence specific anticodon–codon interactions and regulates translation, its efficiency, and fidelity. This phenomenon of nucleoside modification is most remarkable and results in a rich structural diversity of tRNA of which over 93 modifications have been characterized. The variety of post-transcriptional modifications can be classified into two groups according to their complexity. The first group comprises the majority of modified bases, which have simple methylations at the ribose or base moiety that are usually introduced by a single enzymatic reaction. Simple modifications can be found at almost every position of the tRNA molecule with a high density in the tRNA core region, where tertiary interactions between D- and T-arm stabilize the three-dimensional fold, as shown in Figure \(8\). The second group includes complex modifications, whose synthesis requires the sequential activity of several enzymes. Most often these hypermodified nucleosides are found in the anticodon of tRNAs, where they play a direct role in codon recognition and create what is known as the wobble base or wobble position. Panel (A) shows the colored tRNA structure shows the modification frequency of each base. The modification data were taken from the tRNAmodviz database and plotted on the crystal structure of tRNAPhe from S. cerevisiae. Blue-colored bases are rarely modified; red-colored bases are modification hotspots. tRNAs possess two regions with high modification levels—the anticodon loop (especially positions 34 and 37) and the core or elbow region, where D- and T-loop bases interact with each other and stabilize the tertiary fold. For some important positions, the chemical structure of the most frequent modification at this position is shown; Panel (B) shows the three-dimensional structure of pseudouridine at position 55 of tRNAPhe from S. cerevisiae. The additional H-bond donor at N1 interacts with the 5′-adjacent phosphates via a coordinated water molecule. The hydrogen bound to N1 was not resolved in the crystal structure. The ribose shows a stabilizing C3′-endo conformation. PDB: 1EHZ- Panel (C) shows the three-dimensional structure of D16 in the D-arm of tRNAiMet from Schizosaccharomyces pombe. The C5-C6 bond of dihydrouridine is reduced, which leads to a non-planar structure of the base. The ribose takes the less stable C2′ -endo conformation. PDB: 2MN0. A wobble base pair is a pairing between two nucleotides in RNA molecules that does not follow Watson-Crick base pair rules. The four main wobble base pairs are guanine-uracil (G-U), hypoxanthine-uracil (I-U), hypoxanthine-adenine (I-A), and hypoxanthine-cytosine (I-C), as shown in Figure \(9\). To maintain the consistency of nucleic acid nomenclature, “I” is used for hypoxanthine because hypoxanthine is the nucleobase of the inosine nucleotide; nomenclature otherwise follows the names of nucleobases and their corresponding nucleosides (e.g., “G” for both guanine and guanosine – as well as for deoxyguanosine). The thermodynamic stability of a wobble base pair is comparable to that of a Watson-Crick base pair. Wobble base pairs are fundamental in RNA secondary structure and are critical for the proper translation of the genetic code. The wobble base position is usually the first position of the anticodon (read in the 5′ – 3′ direction), which aligns with the 3rd position of the mRNA codon. This helps to explain the degeneracy found within the genetic code as shown in Figure 3 above and Figure \(10\). Degeneracy means that a single tRNA can recognize multiple different codons within mRNA. Panel (A) shows the interaction of the anticodon bases (34–36) of a tRNA with the corresponding bases of the mRNA codons (3, 2, 1). A wobble interaction is possible between codon base 3 and anticodon base 34. The latter is frequently modified and directs the wobble interactions with the third codon base; Panel (B) shows the standard genetic code is illustrated as a simple decoding table, 2-fold degenerate codon boxes are colored yellow, and 4-fold degenerate boxes are blue. Start and stop codons are colored green and red, respectively; Panel (C) shows a stereo image of the well-structured anticodon loop of tRNALys from E. coli. Modifications mnm5s2U34 and t6A37 prevent wrong base pairing inside the 7-nucleotide loop and promote the formation of the conserved U-turn motif. The stacked anticodon bases are located on the same side of the loop. PDB: 1FL8; Panel (D) shows a stereo image of a collapsed and unmodified anticodon loop of tRNATyr from Bacillus subtilis. Here, bases 32 and 38 as well as 33 and 37 interact with each other and the U-turn motif is missing. The anticodon bases are not ordered and are on opposite sides of the loop. PDB: 2LAC. A prominent example is tRNAIle carrying the anticodon UAU. In principle, this anticodon can read codons AUA (for isoleucine) and AUG (for methionine). Yet, it was shown in some instances that tRNAIle with unmodified UAU anticodon exists, but has a strong preference for its cognate AUA codon, while it rarely misreads AUG. In most organisms, however, tRNAIle carries the anticodon CUA. To avoid misreading of the methionine codon by this tRNA, C34 (position 1 of the anticodon) is modified to lysidine (k2C34, with the chemical structure shown in Figure 27.1.10, which restricts codon recognition to only AUA and thereby changes the amino acid identity of the tRNA from methionine to isoleucine. In the archaeal species, Haloarcula marismortui, Methanococcus maripaludis, and Sulfolobus solfataricus, this tRNAIle carries a different modification at C34, fulfilling the same purpose of restricting the interaction to AUA codons. Here, the original cytosine is modified at the C2-oxo position, which is replaced by agmatine (decarboxy-arginine), resulting in agmatidine (C+ or agm2C), as shown in Figure \(11\). A complimentary modification is that of N4-acetylcytosine (ac4C34, whose chemical structure is shown in Figure 27.1.8) in the elongator-tRNAMet of E. coli, which prevents the recognition of the AUA isoleucine codon. In non-plant mitochondria, however, both AUG and AUA codons are read as methionine. Hence, mitochondrial tRNAMet (carrying the anticodon CAU) has to recognize both codon forms. This is achieved by the introduction of 5-formylcytidine (f5C, Figure 11, at position 34, a modification that pairs with both A and U residues at the corresponding codon position 3. The upper part of the image illustrates the systematic abbreviation of RNA modifications with N2,N2,2′-O-trimethylguanosine (m22Gm) as an example and also shows the atom numbering in the purine and pyrimidine rings as well as in the ribose. An abbreviation in front of the base letter describes a base modification, whereas letters after the base stand for ribose alterations. Superscripted numbers specify the position at the base and subscripted numbers indicate the frequency of identical modification at the same position. Abbreviations are as follows: ac—acetyl, acp—aminocarboxypropyl, chm—carboxyhydroxymethyl, cmo—oxyacetic acid, cmnm—carboxymethylaminomethyl, f—formyl, g—glycinyl, gal—galactosyl, hn—hydroxynorvalylcarbamoyl, ho—hydroxy, i—isopentenyl, inm—isopentenylaminomethyl, io—cis-hydroxyisopentenyl, m—methyl, man—mannosyl, mchm—carboxyhydroxymethyl methyl ester, mcm—methoxycarbonylmethyl, mcmo—oxyacetic acid methyl ester, mnm—methylaminomethyl, mo—methoxy, ncm—carbamoylmethyl, nm—aminomethyl, r(p) —5-O-phosphono-b-d-ribofuranosyl, s—thio, se—seleno, t—threonylcarbamoyl, tm—taurinomethyl. The Venn diagram summarizes data collected from the RNA modification database and contains the 93 post-transcriptional modifications that are found in tRNAs. Some examples mentioned throughout the text are shown with their chemical structure. Organisms vary in the number of tRNA genes in their genome. For example, the nematode worm C. elegans, a commonly used model organism in genetics studies, has 29,647 genes in its nuclear genome, of which 620 code for tRNA.The budding yeast Saccharomyces cerevisiae has 275 tRNA genes in its genome. The human genome has approximately 20,848 protein-coding genes, of which there are 497 nuclear genes encoding cytoplasmic tRNA molecules, and 324 tRNA-derived pseudogenes (tRNA genes thought to be no longer functional). Regions in nuclear chromosomes, very similar in sequence to mitochondrial tRNA genes, have also been identified (tRNA-lookalikes). These tRNA-lookalikes are also considered part of the nuclear mitochondrial DNA (genes transferred from the mitochondria to the nucleus). As with all eukaryotes, there are 22 mitochondrial tRNA genes in humans. Mutations in some of these genes have been associated with severe diseases like the MELAS syndrome. Cytoplasmic tRNA genes can be grouped into 49 families according to their anticodon features. These genes are found on all chromosomes, except the 22 and the Y chromosomes. High clustering on 6p is observed (140 tRNA genes), as well as on chromosome 1. Currently, it is unclear why there is so much redundancy within the genome to decode 61 of the 64 possible codons (the other three are stop codons used to terminate translation). Aminoacyl tRNA Synthetases Aminoacyl-tRNA synthetases (aaRSs) are universally distributed enzymes that catalyze the esterification of a tRNA to its cognate amino acid (i.e., the amino acid corresponding to the anticodon triplet of the tRNA according to the genetic code). The product of this reaction, an aminoacyl-tRNA (aa-tRNA), is delivered by elongation factors to the ribosome to take part in protein synthesis. Aminoacyl-tRNA synthetases are named after the aminoacyl-tRNA product generated, as such, methionyl-tRNA synthetase (abbreviated as MetRS) charges tRNAMet with methionine. In eukaryotes, an alternative nomenclature is often employed using the one-letter code of the amino acid (MARS), and a number is added to refer to the cytosolic (MARS1) or the mitochondrial (MARS2) variants. A total of 23 aaRSs have been described so far, one for each of the 20 proteinogenic amino acids (except for lysine, for which there are two) plus pyrrolysyl-tRNA synthetase (PylRS) and phosphoseryl-tRNA synthetase (SepRS), enzymes with a more restricted distribution that are only found in some bacterial and archaeal genomes. It is also worth noting that in eukaryotes the protein synthesis machinery of mitochondria and chloroplasts generally utilize their own, bacterial-like sets of synthetases and tRNAs that are distinct from their cytosolic counterparts. The aminoacyl-tRNA synthetases catalyze a two-step reaction that leads to the esterification of an amino acid to the 3’ end of a tRNA along with the hydrolysis of one molecule of ATP, yielding aminoacyl-tRNA, AMP, and PPi. In the first step, termed amino acid activation, both the amino acid and ATP bind to the catalytic site of the enzyme, triggering a nucleophilic attack of the α-carboxylate oxygen of the amino acid to the α-phosphate group of the ATP, condensing into aminoacyl-adenylate (aa-AMP), which remains bound to the enzyme, and PPi, which is expelled from the active site, as shown in Figure \(12\). Although tRNA is usually not required for this first step, certain synthetases do require the tRNA species for productive amino acid activation. In the second part of the reaction, either the 2′- or 3′-hydroxyl group of the terminal adenine nucleotide attacks the carbonyl carbon of the adenylate, forming aminoacyl-tRNA and AMP (Figure 27.1.12 B). While the two-step aminoacylation reaction is universally conserved, the aaRSs that catalyze it show extensive structural, and in some instances functional, diversity. The 23 known aaRSs can be divided into two major classes based on the architecture of their active sites (Class I and Class II). In class I synthetases, the catalytic domain bears a dinucleotide or Rossman fold (RF) featuring a five-stranded parallel β-sheet connected by α-helices and is usually located at or near the N-terminus of the protein. This RF contains the highly conserved motifs HIGH and KMSKS, separated by a connecting domain termed connective peptide 1 (CP1), as shown in Figure \(13\) (panel A). Class II active site architecture is organized as seven-stranded β-sheets flanked by α-helices and features three motifs that show a lesser degree of conservation than those in class I (panel B). Both classes also exhibit pronounced differences in their modes of substrate binding. Class I aaRSs bind the minor groove of the tRNA acceptor stem (with the exceptions of TrpRS and TyrRS) and aminoacylate the 2’-OH of the ribose of the terminal adenosine, while class II approach tRNA from the major groove and transfer amino acid to the 3’-OH (except PheRS). The mode of ATP binding is also different between both classes, being bound in an extended configuration in class I, while class II binds a bent configuration with the γ-phosphate folding back over the adenine ring. The kinetics of the aminoacylation reaction can also be used as a distinctive mechanistic signature, as aminoacyl-tRNA release is the rate-limiting step for class I enzymes (except for IleRS and some GluRS) while for class II it is the amino acid activation rate instead. Panel (a) shows the E. coliCysRS:tRNACys complex. The CP domain (red) and Rossmann fold catalytic domain (green), stem contact fold (cyan), helical bundle domain (magenta), and anticodon binding domain (orange) of CysRS are shown in a ribbon diagram; Panel (b) A single monomer of the homodimeric E. coli ThrRS:tRNAThr complex. The two N-terminal domains (red), catalytic domain (green), linker (cyan), and anticodon binding domain (orange) of ThrRS are shown in a ribbon diagram. For both structures, the tRNAs are shown in a stick diagram (blue) with a trace of their backbone (yellow). Figure \(14\) shows an interactive iCn3D model of the Class I and II aminoacyl-tRNA synthetases Class I E. Coli Cysteinyl-tRNA synthetase -tRNA(Cys) (1U0B) class II E. coli threonyl-tRNA synthase - tRNA(Thr) (1QF6) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...P4oRWxKvJHFDL6 (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...6aXwfqXGCpD7c6 The color coding is analogous to Figure \(13\) with the tRNA shown in gray. For the Class II E. coli threonyl-tRNA synthase - tRNA(Thr), key conserved amino acids in the active site are shown in CPK-colored sticks and labeled. The anticodon in the tRNA is shown in colored sticks as well. Note the large conformational change in the anticodon loop. The E. coli threonyl-tRNA synthetase is interesting in that it represses the translation of its own mRNA. A Zn2+ ion is involved in binding specificity for the amino acid. It is coordinated by H385, H511, and a water molecule (not shown). R363 interacts with the alpha phosphate, while F379 and R520 align on both sides of the adenine rig. D383 interacts with the amine group of the substrate threonine. To ensure the faithful translation of the genetic message, synthetases must identify and pair particular tRNAs with their cognate amino acid which relies on the proper recognition of both substrates. This can prove extremely challenging for the synthetases as not only have they to discriminate the correct tRNA isoacceptor amongst a set of other tRNAs very similar in structure and chemical composition but also be able to select the cognate amino acid amidst an extremely large pool of similar amino acids, both proteinogenic and non-proteinogenic. The evolutionary pressure to maintain fidelity has driven aaRSs to develop an elevated specificity for their substrates, both the tRNA and the amino acid. In addition, some synthetases have evolved editing activities that specifically target and hydrolyze misactivated amino acids and/or misacylated tRNAs. To date, editing activity has been described in 10 out of the 23 aaRSs. In class I synthetases, this activity is located in the highly conserved CP1 domain, although in some enzymes such as MetRS and LysRS the editing activity resides in the catalytic site. In class II synthetases, however, the editing domains are more idiosyncratic. Editing mechanisms can be divided into two categories, pre- or post-transfer editing, with regard to the editing taking place before or after the transfer of the amino acid to the tRNA, as shown in Figure \(15\). Pre-transfer editing has been described in both class I and class II aaRSs and takes place after aa-AMP synthesis but before the aminoacyl moiety is transferred to the tRNA. Although the tRNA does not participate in the reaction itself, it has been reported that tRNA binding promotes editing activity in some aaRSs and is a requirement in IleRS and LeuRS. Pre-transfer editing can follow two main pathways. The first one is the selective release of the aa-AMP to the cytosol, where the labile phosphoester bond is spontaneously hydrolyzed. The second route involves the enzymatic breakdown of the product and may happen either in the active site or in an independent editing site. Post-transfer editing takes place after the transfer of the amino acid to the tRNA and involves the hydrolysis of the ester bond, in a domain separated from the active site. The specific mechanism of editing is idiosyncratic to each synthetase but in general, once formed the aa-tRNA triggers a conformational change, and the 3’ terminus containing the aa is translocated from the active site to the editing site, sometimes traversing distances as large as 40 Å. As the core of the tRNA remains bound to the enzyme, this translocation often involves a rearrangement of the 3’ terminus to relocate to the editing site. Ribosome Structure The ribosome is a highly conserved molecular machine. In all organisms, it is composed of two unequal subunits, which consist of a distinct set of ribosomal RNA (rRNA) and ribosomal proteins (RPs) that combine to form a large nucleoprotein complex. The ribosome structures in all living organisms harbor three different tRNA binding sites: The A-site, where decoding occurs and the correct aminoacyl-tRNA (aa-tRNA) is selected based on the mRNA codon displayed, the P-site, which carries the peptidyl-tRNA, and the E-site, which binds exclusively deacetylated tRNAs that are exiting the ribosome. Thus, during translation the tRNA moves from the A-site through the P- and E-site, where it leaves the ribosome, as shown in Figure \(16\). The mRNA (shown in purple) is assembled between the small subunit and the large subunit of the ribosome (shown in green). tRNA molecules (shown in red) that are loaded with their cognate amino acid (shown in pink) are transitioned through the A-P-E sites of the ribosome during the elongation phase of translation. Movement of the tRNA molecules also shifts the position of the mRNA causing the next three codon bases to line up in the A-site of the ribosome. The catalytic peptidyl transferase activity occurs when the tRNA molecules are bound in the A- and P-sites, transferring the nascent peptide to the incoming tRNA molecule (Fig. 27.1.15). Ribosomes are ribozymes because the catalytic peptidyl transferase activity that links amino acids together is performed by the rRNA.the complexity of the ribosome structure is reflected in the process of protein synthesis, which can be intersected into three major steps: initiation, elongation, and termination/recycling. Ribosomes are either free-floating in the cytoplasm or they can be associated with the intracellular membranes that make up the rough endoplasmic reticulum (rER). Proteins translated into the rER will often be transported out of the cell or embedded into the plasma membrane. These processes are illustrated in Figure \(17\). Figure \(17\): A ribosome translating a protein that is secreted into the endoplasmic reticulum. Figure from:  Bensaccount Ribosomes from bacteria, archaea, and eukaryotes in the three-domain system resemble each other to a remarkable degree. They differ in their size, much of the rRNA sequence, and the ratio of protein to RNA. Figure \(18\) shows the eukaryotic rRNA from the large subunit of the ribosome with highly conserved nucleotide elements (>90% sequence identity) within all of the domains of life, termed universal CNEs or uCNEs indicated. The differences in sequence and structure between eukaryotes and prokaryotes allow some antibiotics to kill bacteria by inhibiting their ribosomes while leaving human ribosomes unaffected. In all species, more than one ribosome may move along a single mRNA chain at one time (as a polysome), each “reading” its sequence and producing a corresponding protein molecule. In this way, many proteins can be translated from a single mRNA molecule. Within bacteria, translation is also coupled with transcription, as the two processes are not physically separated from one another. This is illustrated in Figure \(19\). In eukaryotic organisms, polysomes form during translation. However, transcription and translation are not coupled, as the processes are separated into the nucleus and cytoplasm, respectively. The mitochondrial ribosomes of eukaryotic cells functionally resemble many features of those in bacteria, reflecting the likely evolutionary origin of mitochondria. Prokaryotic Ribosome Structure Prokaryotic ribosomes have a mass of about 2500 kDa and a size of 70S (or Svedberg units: A Svedberg unit is a measure of the sedimentation rate in a centrifuge and thus is representative of size). A complete ribosome (70S) can be dissociated into a large subunit (50S) and a small subunit (30S), as shown in Figure \(20\). The small subunit is formed by the interactions of 21 different proteins and a 16S RNA molecule, whereas the large subunit contains 34 different proteins and two RNA molecules, a 23S, and a 5S species. The rate-limiting step in protein synthesis is the formation of the 70S initiation complex which will be discussed in detail in the next section. Figure \(21\) shows an interactive iCn3D model of the E. Coli ribosome (7K00). The RNA is shown in a faint trace backbone. The proteins are shown as cartoons with different colors. Note the large number of Mg2+ ions. The structure was determined using cryo-EM at high resolution. Some ribosomal proteins have isopeptide bonds (using side chain amine and carboxyl groups) as well as some thioamide backbone replacements for the usual amide links. Eukaryotic Ribosome Structure Eukaryotic ribosomes are larger than their prokaryotic counterparts at approximately 80S (although there is some modest variation between eukaryotic species). Human cytosolic ribosomes are composed of a large subunit (60S) that contains the 28S, 5.8S, and 5S rRNAs and 47 ribosomal proteins (RPs), and a small subunit (40S) that contains the 18S rRNA and 33 RPs. The assembly of eukaryotic ribosomal subunits starts in the nucleolus, where RNA polymerase I transcribes the major rRNA precursor (a 45S pre-RNA), from which, after processing and removal of the external and internal transcribed spacers (ETS and ITS), the mature rRNAs are generated, as shown in Figure \(22\). The pre-RNA is modified during transcription by small nucleolar ribonucleoproteins (snoRNPs), processed by RNA nucleases, and assembled with numerous RPs. After processing the rRNA precursor, the pre-40S and pre-60S subunits follow separate biogenesis routes. Here we will describe the assembly of the 60S subunit in more detail. Although the exact assembly of the 60S subunit is not currently known, a model has been postulated that suggests that in the nucleolus, after circularization of rRNA domains, early 60S assembly is carried out sequentially, as shown in Figure \(23\). As the transcription of the pre-rRNA proceeds, the rRNA quickly develops a core secondary structure that promotes the interaction of key Assembly Factors (AFs) and RPs before transcriptional termination. Specifically, during this time, the 5.8S portion, ITS2, domains I and II, and partially domain VI are folded in the earliest observed intermediate (state A in Fig. 23). Thus, it appears that the solvent-exposed back side of the large subunit forms like an exoskeleton and construction proceeds by formation of the exit tunnel. This model agrees with a previously suggested model of hierarchical folding based on RP depletion phenotypes. The peptidyl transferase center (PET) is predicted to be one of the later folding steps in the process (state F in Fig. Fig 23) Although the very late-folding peptidyl transferase center is the evolutionarily oldest part of the ribosome, it is likely that folding the solvent side first brings the advantage of providing a stable scaffold for the developing 60S subunit. The folding and assembly of the 40S subunit follow a similar pattern. The two subunits remain unattached until activated in the cytoplasm through the binding of a mRNA transcript with the small subunit. This begins the formation of the initiation complex that will mark the start of the transcriptional process. Assembly of RPs and AFs to the nascent 35S rRNA precursor starts co-transcriptionally. Very early, the pre-rRNA is circularized as domain VI binds to domains I and II and the 5.8S portion of the precursor rRNA. The formation of the PET (displayed here as a black circle) starts with this circularization. Its maturation progresses as rRNA domains fold following this order: VI, V, III, and IV. Full assembly of the PET is only achieved when domain V is completely folded as observed in state F. After that, only a few additional steps need to occur before the particles are exported to the cytoplasm, where they undergo final maturation.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/26%3A_Protein_Metabolism/26.01%3A_The_Genetic_Code.txt

Search Fundamentals of Biochemistry Prokaryotic Initiation The small subunit of the ribosome (the 30S) interprets the genetic information by selecting aminoacyl-tRNAs cognate to the mRNA codons in the decoding center. The large subunit (the 50S) carries the catalytic peptidyl transferase center where amino acids are polymerized into a protein. Small and large subunits unite together at the start codon of a gene to form the 70S ribosome and dissociate again at the stop codon upon completing the synthesis of the encoded protein. This process consists of three phases: initiation, elongation, and termination. In this section, we will focus on the initiation of translation. In bacteria, the initiation phase of protein synthesis involves a limited number of “actors”. Aside from the two ribosomal subunits, key roles are played by the initiator tRNAfmet, the translation initiation region (TIR) of the mRNA, and three protein factors – the initiation factors (IFs) IF1, IF2, and IF3 – that ensure speed and accuracy to the overall process. The initiator tRNAfmet contains a methionine residue that has been enzymatically modified to contain an N-terminal formyl group, as shown in Figure \(1\). fMet is only used for the initiation of protein synthesis and is thus found only at the N-terminus of the protein. Unmodified methionine is used during the rest translation. Once protein synthesis is completed, the formyl group on methionine may be removed by peptide deformylase, and on occasion, the entire methionine residue can be further removed by the enzyme methionine aminopeptidase. The TIR sequence within the mRNA contains the start codon and usually an upstream untranslated region that interacts with the small subunit of the ribosome. The bacterial cell produces and expresses a plethora of different mRNAs with different TIR sequences and structures; the efficiency by which these individual transcripts are translated depends not only upon their abundance and stability but also upon the nature of TIR. Thus, unlike the other aforementioned actors that represent constants, the mRNA TIRs represent essentially the only variable in the process of mRNA initiation site selection and can affect translation efficiency. Although the triplet AUG is by far the most frequent initiation codon found in TIRs, other initiation triplets (i.e., GUG, UUG, AUU, AUC, and AUA) are found in bacteria and the central U is the only universally conserved base of the start codon. Among the aforementioned triplets, those having a 3′-G (i.e., AUG, GUG, and UUG) are recognized equivalently and most efficiently by IF3 during the initiation complex formation. Another important characteristic of a large number of bacterial mRNA TIRs is the presence of a Shine–Dalgarno (or SD) sequence that is complementary to the 3′ end sequence of 16S rRNA (the anti-SD sequence or aSD). The SD sequence, when present, is usually at an optimal distance of 4–9 nucleotides upstream of the initiation codon, as shown in Figure \(2\). While the SD sequence plays an important role in the efficient translation of many mRNA transcripts, it is not essential. Many other mRNA sequences fully lack an SD sequence but are still efficiently transcribed. Thus, the SD sequence is only one example of TIR mechanisms that can play an important role in mRNA binding with the small subunit of the ribosome. Prokaryotic mRNA sequences often share a highly conserved sequence upstream of the start codon known as the Shine-Dalgarno sequence. This consensus sequence is complimentary to the 3′-end of the 16S rRNA sequence in the small subunit of the ribosome. It is an important feature for the binding and docking of many mRNA molecules with the small ribosomal subunit during transcription initiation. The three protein initiation factors, IF1, IF2, and IF3, determine the kinetics and fidelity of the overall initiation process. The three IFs are bound, one copy each, to specific sites of the 30S subunit where they assist with the formation of the initiation complex and assembly of the 70S ribosome. As noted above, the initiator tRNA is first aminoacylated with methionine whose α-NH2 group is eventually blocked by a specific formyl transferase (TMF) to produce a tRNAfmet molecule. This modification prevents interaction with the elongation factor EF-Tu (which we will see plays an important role in the elongation phase of translation, but not the initiation phase!) Blocking EF-Tu binding ensures instead the recognition and binding of tRNAfmet by initiation factor IF2, effectively docking it with the 30S subunit. Furthermore, tRNAfmet binds with high affinity to the ribosomal P-site, unlike all other aminoacyl-tRNAs that bind to the A-site in a ternary complex with EF-Tu and GTP (details will be presented in the next section). In the P-site, the initiator tRNA must be recognized as correct by the other initiation factors IF3 and IF1. To form the 30S initiation complex, IF3, and IF2 are the first factors to bind to the 30S subunit forming an unstable 30S-IF3-IF2 complex, as shown in Figure \(3\) (panel A). The binding of IF1 causes a conformational change in the 30S subunit stabilizing the complex and allowing the recruitment of the tRNAfmet by IF2. Notably, IF1 binds in the A site of the 30S subunit, where it contacts ribosomal protein S12. Recruitment of the tRNAfmet can also stabilize the mRNA interactions with the 30S subunit through the formation of hydrogen bonds between the codon of the mRNA and the anticodon of the tRNAfmet. Note that the binding of mRNA to the 30S subunit is IF-independent and can take place at any time during the 30S assembly process. Two potential routes of mRNA association are shown Figure \(3\), panel A, where the mRNA is assembled either prior to or after tRNAfmet recruitment. Step 1: a vacant 30S ribosomal subunit binds IF3 and IF2. Step 2: IF1 binds to the 30S subunit in the presence of both IF3 and IF2. Steps 3 and 3′: in the presence of all three factors tRNAfmet is recruited. Steps 4 and 4′:the mRNA is bound with different on and off rates depending on its TIR structure; mRNAs with strong secondary structures are bound more slowly than those having little or no secondary structure. Step 5: mRNAs containing secondary structures must be unfolded in a process that is facilitated by IF2 bound to GTP and antagonized by IF3. Step 6: the isomerization of the 30S pre-IC allows the P-site codon–anticodon interaction to yield a more stable 30SIC from which mRNA and fMet-tRNA are more stably bound. Step 7: a 30SIC, containing IF1, IF2·GTP, IF3 and mRNA whose initiation triplet is P-site decoded by fMet-tRNA, is docked by a 50S subunit. Step 8: upon contact with the 50S subunit, the GTPase function of IF2 is activated and GTP is rapidly hydrolyzed leaving GDP+Pi bound to IF2. Step 9: this reversible conformational transition represents the last kinetic checkpoint of translation initiation fidelity by IF3 and IF1, as IF3 and IF1 dissociate from the complex. Step 10: The first-order isomerization of the IF2-GDP structure causes a shift in the ribosome structure that represents the rate-limiting step in 70SIC formation. Step 11: Pi is dissociated from IF2·GDP. Step 12: IF2 leaves the ribosome (or moves away from the A-site) clearing the way for EF-Tu binding. Step 13: the EF-Tu·GTP·aminoacyl-tRNA complex binds to the 70SIC and through a number of steps (not represented here) delivers to the ribosomal A-site the aminoacyl-tRNA encoded by the second mRNA codon. Step 14:the tRNAfMet bound in the P-site of the peptidyl transferase center donates its formyl-methionine to the A-site-bound aminoacyl-tRNA to yield the initiation dipeptide fMet-aa. Initiation is then complete and the elongation phase can begin. Following the recruitment of the mRNA and the tRNAfmet to the 30S initiation complex loaded with the IF2, IF3 and IF1 initiation factors, the 50S subunit is very rapidly docked to yield an initially unstable 70S initiation complex (Fig 27.2.3 b). It should be noted that the IF2 protein is a GTP hydrolase enzyme and, as such, binds with the cofactor GTP prior to the recruitment of the 50S subunit. Contact between the IF2 GTPase activating center with the 50S subunit causes the rapid hydrolysis of GTP to GDP + Pi. The formation of the 70S complex causes the dissociation of the initiation factors. IF2 is the last factor to be dissociated, leaving the ribosome after having positioned tRNAfMet in the P-site of the 70S initiation complex. It must be placed in the correct orientation to facilitate peptide bond formation. GDP and Pi also dissociate from the complex with the removal of IF2. The elongation factor, EF-G is then free to chaperone the first tRNA into the A-site and the first peptide bond is formed (Step 13 of Fig. 27.2.3 b). This marks the beginning of the elongation phase of protein synthesis. Eukaryotic Initiation Eukaryotic translation initiation is more complex than prokaryotic systems and requires the actions of at least 11 eukaryotic initiation factors (eIFs), plus additional auxiliary factors (Table \(1\)). We will not cover the action of all these eIFs in detail here, but rather focus a few key steps as outlined in Figure \(4\). Table \(1\): Comparison of Prokaryotic and Eukaryotic Translation Initiation Factors First, the initiator tRNAi is recruited to the small ribosomal subunit (40S) to form a ternary complex with the GTP-bound eukaryotic initiation factor 2 (eIF2). Formation of this 43S pre-initiation complex is strongly enhanced by additional factors, such as eIF3. eIF3 also interacts with the eIF4F complex, which consists of three other initiation factors: eIF4A, eIF4E, and eIF4G. eIF4G is a scaffolding protein that directly associates with both eIF3 and the other two components. eIF4E is the 5′-cap-binding protein. Binding of the mRNA cap by eIF4E is often considered the rate-limiting step of cap-dependent initiation, and the concentration of eIF4E is a regulatory nexus of translational control. Certain viruses cleave a portion of eIF4G that binds eIF4E, thus preventing cap-dependent translation to hijack the host machinery in favor of the viral (cap-independent) messages. eIF4A is an ATP-dependent RNA helicase that aids the ribosome by resolving certain secondary structures formed along the mRNA transcript. The poly(A)-binding protein (PABP) also associates with the eIF4F complex via eIF4G, and binds the poly-A tail of most eukaryotic mRNA molecules. This protein has been implicated in playing a role in circularization of the mRNA during translation.The 43S preinitiation complex accompanied by the protein factors moves along the mRNA chain toward its 3′-end, in a process known as ‘scanning’, to reach the start codon (typically AUG). After recognition of the start codon, the large ribosomal subunit (60S) assembles to form the 80S initiation complex, which is ready for elongation. Alternatively, under distinct conditions or on certain transcripts internal initiation can occur in a cap-independent manner at so called internal ribosome entry sites (IRES). Eukaroytic translation initiation is shown in Figure \(4\). This is a simplified diagram of eukaryotic translation initiation detailing some of the eIFs involved in the process. eIF2 is critical for recruiting the initiation tRNAi to the 40S subunit. eIF3 enhances the activity of eIF2 and also promotes the binding of the 43S pre-initiation complex with the mRNA. eIF3 binds with the mRNA through the interaction of the eIF4 factors and causes the scanning of the pre-initiation complex down the mRNA to locate the start codon (usually AUG). Poly A Binding Proteins (PABPs) bind with the polyA tail sequence of the mRNA and also interact with the eIF4 factors causing the circularization of the mRNA. As seen in prokaryotic systems with the favored Shine Dalgarno sequence upstream of the start codon within the mRNA sequence, there are also preferred nucleotide sequences within the local vicinity of the start codon in eukaryotic mRNAs, as well. In eukaryotic mRNA, this is known as the Kozak sequence (Figure \(5\)). The sequence was originally defined as 5′-`(gcc)gccRccAUGG-3` where: 1. The underlined nucleotides indicate the translation start codon, coding for Methionine. 2. upper-case letters indicate highly conserved bases, i.e. the ‘AUGG’ sequence is constant or rarely, if ever, changes. 3. ‘R’ indicates that a purine (adenine or guanine) is always observed at this position (with adenine being more frequent according to Kozak rules) 4. a lower-case letter denotes the most common base at a position where the base can nevertheless vary 5. the sequence in parentheses (gcc) is of uncertain significance. The AUG is the initiation codon encoding a methionine amino acid at the N-terminus of the protein. (Rarely, GUG is used as an initiation codon, but methionine is still the first amino acid as it is the met-tRNA in the initiation complex that binds to the mRNA). Variation within the Kozak sequence alters the “strength” of the translational start site. Kozak sequence strength refers to the favor ability of initiation, affecting how much protein is synthesized from a given mRNA. This is shown in Figure \(5\). The Elongation Phase of Translation Both prokaryotic and eukaryotic elongation phases of transcription utilize similar elongation factors during the process. Table \(2\) provides a summary of their functions. Table \(2\): Comparison of Prokaryotic and Eukaryotic Translation Elongation Factors Prokaryotic Elongation The prokaryotic elongation phase of transcription requires the activity of three primary elongation factors (EFs), EF-Tu, EF-Ts, and EF-G. During elongation, aminoacyl-tRNAs are delivered to the ribosome in the form of a ternary complex: the tRNA, a translational GTPase (in bacteria: EF-Tu or SelB), and a GTP molecule, as shown in Figure \(6\). The tRNA decodes the information on the mRNA by forming hydrogen bonds (H-bonds) between codon and anticodon nucleobases. Remarkably, the free-energy difference between correct (cognate) and incorrect (near-cognate, non-cognate) base pairing alone does not explain the very high fidelity of decoding. Rather, high fidelity is achieved by a two-step decoding process: initial selection leading to GTPase activation and proofreading. In addition to the free-energy difference, kinetic effects contribute to the discrimination. The GTP hydrolysis rate is increased and tRNA rejection rate is decreased by the recognition of the correct codon. Small-subunit nucleotides A1492 and A1493 adopt a flipped-out conformation in the presence of a tRNA and, in this conformation, the tRNA anticodon hydrogen bonds with the codon of the mRNA forming a mini-helix structure as shown in Figure \(7\). The flipped out nucleotides A1492, A1493 along with G530 were found to shield the codon–anticodon base pairs from solvent. This shielding of near-cognate base pairs from the solvent is incomplete causing an increase in the free-energy difference between near-cognate and cognate base pairs and more flexibility within the docking region. This reduces the strength of hydrogen bonding between a non-cognate tRNA and causes the inappropriate tRNA to leave the A-site before peptide bond formation can occur. This increases the fidelity and discrimination of tRNA selection, such that only the correct cognate tRNA is incorporated into the A-site. Interestingly aminoglycosides, a class of antibiotics, bind to the decoding center and lock nucleotides A1492/A1493 in the flipped-out conformation as shown in Figure \(8\). In this way aminoglycosides promote the accommodation of near-cognate, thus wrong, tRNAs into proteins during synthesis causing wide-spread mutagenesis. This is toxic to the bacteria and leads to bacterial cell death. After GTP hydrolysis, the GTPase EF-Tu dissociates from the tRNA. At this point, the EF-Tu is tightly bound with a molecule of GDP and cannot release GDP on its own to be recycled for a second round of tRNA chaperoning. The recharging of EF-Tu is executed by the Elongation Factor Thermo stable (EF-Ts), as shown in Figure \(9\). The binding of EF-Ts with EF-Tu-GDP causes a conformational change in EF-Tu that allows the release of GDP. The binding of a new molecule of GTP with the EF-Tu protein causes the dissociation of EF-Ts and fully recharges EF-Tu. Dissociation of EF-Tu from the ribosome allows the tRNA to move into to the peptidyl transferase center (A-site) on the large subunit. At the core of ribosomal translation is the catalysis of peptide bond formation, as shown in Figure \(10\). The current reaction models point to a substrate assisted mechanism. Simulations indicate that the transition state forms due to extensive hydrogen bonding with water molecules and the surrounding rRNA bases and that the C-O bond cleavage takes place after C-N bond formation. Peptide bond formation results in the transition of the amino acid docked on the P-site tRNA to the nascent growing peptide that is now held on the tRNA in the A-site. Note that this mechanism causes the nascent growing peptide to always grow in the N- to C- direction. Once the peptide bond is formed, the ribosome needs to translocate down the mRNA to make the next mRNA codon available within the A-site. This also requires the shifting of the tRNA molecules, such that the tRNA in the A-site (which is now tethered to the nascent peptide) shifts to the P-site. The P-site tRNA (which is now empty) shifts to the E-site, and if there was an empty tRNA in the E-site, it will shift to exit the ribosome. Shifting the tRNAs and mRNAs within the ribosome core requires the action of the EF-G elongation factor, as shown in Figure \(11\). Figure \(12\) shows an interactive iCn3D model of the eukaryotic 80S ribosome with bound mRNA and tRNAs (6GX3) . (Very long load time) color coding as follows: • gray: protein tube • coiled coils: RNA trace • black spheres: mRNA • dark blue spheres: ap/P-site tRNA • cyan spheres: pe/E-site-tRNA • green spheres: Mg2+ EF-G is a GTP hydrolase protein that binds to the A-site of the ribosome. The EF-G protein has high flexibility that enables it to act as a hinge. Folding of EF-G is dependent on GTP hydrolysis. Thus, when binding to the ribosome, the fast hydrolysis of GTP acts as a power stroke folding the EF-G protein and causing a conformation shift in the ribosome that enables the translocation of the tRNA residues and the mRNA. Translocation of tRNAs is accompanied by large-scale collective motions of the ribosome: relative rotation of ribosomal subunits and L1-stalk motion, as shown in Figure \(13\). The L1 stalk, which is a flexible part of the large subunit, is in contact and moves along with the tRNA from the P to the E site. Once in the EF-G-GDP form, the factor quickly dissociates from the ribosome, opening up the A-site for the recruitment of the next aa-tRNA molecule. The elongation cycle will continue to be repeated until a termination codon is reached. Eukaryotic Elongation The elongation phase in eukaryotic translation is very similar to prokaryotic elongation. Essentially, the mRNA is decoded by the ribosome in a process that requires the selection of each aminoacyl-transfer RNA (aa-tRNA), which is dictated by the mRNA codon in the ribosome acceptor (A) site, peptide bond formation and movement of both tRNAs and the mRNA through the ribosome, as shown in Figure \(14\). A new amino acid is incorporated into a nascent peptide at a rate of approximately one every sixth of a second. The first step of this process requires guanosine triphosphate (GTP)-bound eukaryotic elongation factor 1A (eEF1α) to recruit an aa-tRNA to the aminoacyl (A) site, which has an anticodon loop cognate to the codon sequence of the mRNA. The anticodon of this sampling tRNA does not initially base-pair with the A-site codon. Instead, the tRNA dynamically remodels to generate a codon-anticodon helix, which stabilizes the binding of the tRNA-eEF1α-GTP complex to the ribosome A site. This helical structure is energetically favorable for cognate or correct pairing, and so discriminates between the non-cognate or unpaired and single mismatched or near-cognate species. This is important for the accuracy of decoding since it provides a mechanism to reject a non-cognate tRNA that carries an inappropriate amino acid. The pairing of the tRNA and codon induces GTP hydrolysis by eEF1α, which is then evicted from the A site. In parallel with this process, the ribosome undergoes a conformational change that stimulates contact between the 3′ end of the aa-tRNA in the A site and the tRNA carrying the polypeptide chain in the peptidyl (P) site. The shift in position of the two tRNAs [A to the P site and P to the exit (E) site] results in ribosome-catalyzed peptide bond formation and the transfer of the polypeptide to the aa-tRNA, thus extending the polypeptide by one amino acid. The second stage of the elongation cycle requires a GTPase, eukaryotic elongation factor 2 (eEF2), which enters the A-site and, through the hydrolysis of GTP, induces a change in the ribosome conformation. This stimulates ribosome translocation to allow the next aa-tRNA to enter the A-site, thus starting a new cycle of elongation. This schematic represents the four basic steps of eukaryotic translation elongation. The ribosome contains three tRNA-binding sites: the aminoacyl (A), peptidyl (P) and exit (E) sites. In the first step of peptide elongation, the tRNA, which is in a complex with eIF1 and GTP and contains the cognate anticodon to the mRNA coding sequence, enters the A site. Recognition of the tRNA leads to the hydrolysis of GTP and eviction of eEF1 from the A site. In parallel, the deacylated tRNA in the E site is ejected. The A site and the P site tRNAs interact, which allows ribosome-catalyzed peptide bond formation to take place. This involves the transfer of the polypeptide to the aa-tRNA, thus extending the nascent polypeptide by one amino acid. eIF5A allosterically assists in the formation of certain peptide bonds, e.g. proline-proline. eEF2 then enters the A site and, through the hydrolysis of GTP, induces a change in the ribosome conformation and stimulates translocation. The ribosome is then in a correct conformation to accept the next aa-tRNA and commence another cycle of elongation. The Ribosome as a Ribozyme Protein synthesis from a mRNA template occurs on a ribosome, a nanomachine composed of proteins and ribosomal RNAs (rRNA). Peptide bond formation occurs when another tRNA-amino acid molecule binds to an adjacent codon on mRNA. The tRNA has a cloverleaf tertiary structure with some intrastranded H-bonded secondary structure. The last three nucleotides at the 3' end of the tRNA are CpCpA. The amino acid is esterified to the terminal 3'OH of the terminal A by a protein enzyme, aminoacyl-tRNA synthetase. Covalent amide bond formation between the second amino acid to the first, forming a dipeptide, occurs at the peptidyl transferase center, located on the larger ribosomal subunit (50S and 60S in bacteria and eukaryotes, respectively). The ribosome ratchets down the mRNA so the dipeptide-tRNA is now at the the P or Peptide site, awaiting a new tRNA-amino acid at the A or Amino site. Figure \(15\): below shows a schematic of the ribosome with bound mRNA on the 30S subunit and tRNAs covalently attached to amino acid (or the growing peptide) at the A and P site, respectively. A likely mechanism (derived from crystal structures with bound substrates and transition state analogs) for the formation of the amide bond between a growing peptide on the P-site tRNA and the amino acid on the A-site tRNA is shown in Figure \(16\). Catalysis does not involve any of the ribosomal proteins (not shown) since none is close enough to the peptidyl transferase center to provide amino acids that could participate in general acid/base catalysis, for example. Hence the rRNA must act as the enzyme (i.e. it is a ribozyme). Initially it was thought that a proximal adenosine with a perturbed pKa could, at physiological pH, be protonated/deprotonated and hence act as a general acid/base in the reaction. However, none was found. The most likely mechanism to stabilize the oxyanion transition state at the electrophilic carbon attack site is precisely located water, which is positioned at the oxyanion hole by H-bonds to uracil 2584 on the rRNA. The cleavage mechanism involves the concerted proton shuffle shown below. In this mechanism, the substrate (Peptide-tRNA) assists its own cleavage in that the 2'OH is in position to initiate the protein shuttle mechanism. (A similar mechanism might occur to facilitate hydrolysis of the fully elongated protein from the P-site tRNA.) Of course all of this requires perfect positioning of the substrates and isn't that what enzymes do best? The main mechanisms for the catalysis of peptide bond formation by the ribosome (as a ribozyme) are intramolecular catalysis and transition state stabilization by the appropriately positioned water molecule. Translation Termination Prokaryotic Termination Termination of bacterial protein synthesis occurs when a stop codon is presented in the ribosomal A-site and is recognized by a class I release factor, RF1 or RF2. These release factors (RFs) have different but overlapping specificities, where RF1 reads UAA and UAG and RF2 reads UAA and UGA, with strong discrimination against sense codons. The RFs are multi-domain proteins, where binding and stop codon recognition by domain 2 at the decoding site causes the universally conserved GGQ motif of domain 3 to insert into the A-site of the PTC, some 80 Å away from the decoding site. This event triggers hydrolysis of the peptidyl-tRNA bond in the P-site of the PTC, and the nascent peptide chain can then be released via the ribosomal exit tunnel, as shown in Figure \(14\). After peptide release, RF1 and RF2 dissociate from the post-termination complex. The dissociation is accelerated by a class II release factor called RF3, which functions as a translational GTPase that binds and hydrolyses GTP in the course of termination. While RF3 increases the efficiency of peptide hydrolysis, it is not an essential protein for the process. In gene knockout studies, RF3 is dispensable for growth of Escherichia coli, and its expression is not conserved in all bacterial lineages. For example, RF3 is not present in the thermophilic model organisms of the Thermus and Thermatoga genera and in infectious Chlamydiales and Spirochaetae. This means that both RF1 and RF2 are capable of performing a complete round of termination independently of RF3 or that other GTPases from the elongation or initiation phases of translation can compensate for the action of RF3. The release factors RF1 and RF2 acquire an open conformation (Figure \(17\) on the 70S ribosome, which is distinctly different from the closed conformation observed in crystal structures of free RFs. The conformational equilibrium of the free RFs in solution shows that this open conformation is dominating at about 80%. During peptide hydrolysis, the RF factors cause rotational and conformational changes within the ribosome that allow the binding of a ribosome recycling factor (RRF) and the EF-G GTPase, which leads to the dissociation of the large subunit from the small subunit and the release of the mRNA, as shown in Figure \(18\). When a stop codon enters the A-site of the ribosome RF1 or RF2 enter the A-site and bind with the mRNA. This leads to the hydrolysis of the protein and release through the exit tunnel. Binding of RF3 and GTP hydrolysis causes the dissociation of the RF factors and conformational change of the ribosome structure. Subse Eukaryotic Termination In eukaryotes and archaea, on the other hand, a single omnipotent RF reads all three stop codons. Although the mechanism of translation termination is basically the same, there is neither sequence nor structural homology between the bacterial RFs and the eukaryotic eRF1, apart from the universally conserved GGQ motif which is required for peptide hydrolysis from the tRNA. the eRF3 GTPase coordinates the release of eRF1 following hydrolysis. In Archaea, there is no eRF3 homolog, instead the aEF1A protein mediates this function. The process of eukaryotic ribosomal disassembly and recycling is currently not well understood, but appears to involve an ABC type ATPase called ABCE1. Mitochondria have independent RFs that can recognize standard and non-standard stop codons, and are more homologous with bacterial systems of ribosomal recycling and disassembly. Summary of Translation An overall summary of prokaryotic translation is given in Figure \(19\). Left panel: Structure of the bacterial ribosome in complex with EF-Tu (PDB 5AFI). Right Panel: Scheme of the bacterial translation cycle. 30S: small subunit; 50S: large subunit; IF1, IF2, IF3: initiation factors; fM-tRNA: N-formylmethionine tRNA; aa-tRNA: aminoacyl tRNA; EF-Tu, EF-G: elongation factors; RF1, RF2, RF3: release factors; RRF: ribosome recycling factor; green trace: nascent protein. The question mark stands for a stop codon recognition.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/26%3A_Protein_Metabolism/26.02%3A_Protein_Synthesis.txt

Search Fundamentals of Biochemistry Regulation of Translation Heterogeneity of Ribosome Structure Over the years, many studies performed in eukaryotes presented evidence that ribosomes can vary in their protein and rRNA complement between different cell types and developmental states. These observations culminated in the postulation of the ‘ribosome filter hypothesis’ by Mauro and Edelman in the year 2002. The authors propose that the ribosome composition functions as a translation determination factor. Depending on the RPs and rRNA sequences represented in the respective ribosome, the complex acts like a filter that selects for specific mRNAs and hence modulates translation, as shown in Figure \(1\). RP heterogeneity can arise from differential expression of paralogs/homologs of RP proteins within different cell types or occur due to differential post-translational modifications of RPs, such as phosphorylation. The protein-to-rRNA ratio may also slightly vary within ribosomal composition affecting translation efficiency and selectivity. RNA genes are also present in multiple copies throughout the genomes of organisms from all domains of life. For example, the bacteria Streptomyces coelicolor harbors six copies of divergent large subunit (LSU) rRNA genes that constitute at least five different LSU rRNA species in a cell. These genes were shown to be differentially transcribed during the morphological development of the organism. Similarly, B. subtilis harbors ten rRNA operons and their reduction to one copy increased the doubling time as well as the sporulation frequency and the motility of the resulting mutant. Modification of the rRNA also provides another avenue of ribosomal heterogeneity. Similar to tRNA, rRNA residues can be chemically modified and commonly have 2-OH methylation. The conversion of uridine to pseudouridine is also quite common. In eukaryotes, the modifications are facilitated by snoRNAs and their tissue-specific expression might be a source for ribosome specialization. In light of the increasing evidence, ribosome heterogeneity, though still far from being entirely understood, proves to be an integral mechanism to modulate and fine-tune protein synthesis in response to environmental signals in all organisms. Effects of Sequence and Secondary Structure in mRNA The amount of protein produced from any given mRNA (i.e., the translational output) is influenced by multiple factors specified by the primary nucleotide sequence. These factors include GC content, codon usage, codon pairs, and secondary structure. For example, 5’UTR sequences in the mRNA may interact with small miRNAs and lead to RNA interference. miRNA interactions may also target mRNA for degradation(Figure \(2\)). This process is aided by protein chaperones called argonautes. This antisense-based process involves steps that first process the miRNA so that it can base-pair with a region of its target mRNAs. Once the base pairing occurs, other proteins direct the mRNA to be destroyed by nucleases. Fire and Mello were awarded the 2006 Nobel Prize in Physiology or Medicine for this discovery. • Step (1) shows how Exportin-5 transports a hairpin primary micro RNA (pri-miRNA) out of the nucleus and into the cytoplasm. • Step (2) shows how Dicer (not shown) trims the pri-miRNA and removes the hairpin loop. A group of proteins, known as Argonautes, form a miRNA/protein complex. • Steps (3,4) show how miRNA/protein complex hydrogen bonds with mRNA based on complimentary sequence homology, and blocks translation. • Step (5) shows the miRNA/protein complex binding speeds up the breakdown of the polyA tail of the mRNA, causing the mRNA to be degraded sooner. Effects of the Nascent Peptide on Ribosome Efficiency Since the Peptidyl Transferase Center (PTC) is buried within the large subunit, during translation the nascent peptide chain (NC) exits through a 100 Å-long tunnel (Figure \(3\)). The exit tunnel plays an active role in protein synthesis. Certain peptide sequences specifically interact with tunnel walls and induce ribosome stalling. Furthermore, the exit tunnel is a binding site for a clinically important class of antibiotics known as the macrolides. When synthesizing proteins containing proline stretches (i.e. several prolines in a row), ribosomes become stalled. Stalling is alleviated by a specialized elongation factor, EF-P in bacteria. Recently, cryo-EM structures of a ribosome stalled by a proline stretch with and without EF-P were resolved. In simulations of the PTC region, elongation factor P (EF-P) was observed to stabilize the P-site tRNA in a conformation compatible with peptide bond formation, while in the absence of EF-P, the P-site tRNA moved away from the A-site tRNA. The exit tunnel can accommodate 30–60 AAs, depending on the level of NC compaction. The rate of translation of about 4–22 AA per second in bacteria provides the NC with sufficient time to explore its conformational space and to start folding when still bound to the ribosome-tRNA complex. Proteasome The 26S proteasome is the central element of proteostasis regulation in eukaryotic cells. It is required for the degradation of protein factors in multiple cellular pathways and it plays a fundamental role in cell stability. The 26S proteasome has a structural configuration that confines the proteolytic active sites in a location unreachable for native and functional proteins, thus preventing uncontrolled degradation. The proteolytic active sites are found in the interior of a barrel-shaped core particle (CP or 20S). The entrances of the tunnel, placed at the distal ends of the barrel, are commonly occupied by the regulatory particle (RP or 19S), a sophisticated protein assembly that acts as a substrate processing machine. The regulatory particle has the important role of receiving, deubiquitinating, unfolding, and translocating substrates to the CP and it adopts different configurations depending on the activity states they exhibit. This process typically requires ATP hydrolysis. Moreover, conformationally distinct proteasomes may show different subcellular distributions depending on functional requirements in each cell type and environmental situation. Proteasomes are distributed throughout the cell, detected in the cytoplasm and the nucleus, and they can localize to hotspots in distinct intracellular regions or specific sites with high protein metabolism or with specific protein degradation requirements. The core of the proteasome consists of a symmetrical cylinder-shaped structure composed of four stacked rings, each containing 7 different subunits and is called the 20S proteasome, as shown in Figure \(4\). The two outer rings are each composed of seven α-subunits (α1-α7 or PSMA1-7). During proteasome assembly, the α-rings serve as the backbone for the incorporation of β-subunits, followed by the dimerization of two half proteasomes. In mature proteasomes, the α-rings regulate substrate entrance since the α-subunits have hydrophobic loops that close the 20S barrel to prevent the random entry of substrates. The 20S core of the proteasome consists of 4 stacked rings. The outer rings contain seven α-subunits (white) while the inner rings contain seven β-subunits (purple). The catalytic subunits, β1, β2, and β5, are depicted in shades of blue. Gate opening of the 20S core occurs via capping by proteasome activators such as the 19S cap or PA28. The 19S cap is the most abundant activator and it forms the 26S proteasome together with the 20S core. Different cells have different caps. For example, interferoIFN-γ stimulation induces de novo formation of immunoproteasomes, which contains the immune subunits β1i (LMP2), β5i (LMP7), and β2i (MECL-1) (shades of red), as well as proteasome activation by PA28αβ (shades of green). Proteasomes in neural tissue are discussed below. In general, protein entry can only be established after gate opening by proteasome activators (PAs) such as the 19S cap, after which substrates can enter the interior of the 20S core for degradation. The inner two rings of the 20S barrel consist of the subunits β1-β7 (PSMB1-7). Each β-ring contains 3 catalytic subunits; termed β1, β2, and β5. In mature 20S complexes, the pro-peptides of these catalytic β-subunits are auto-catalytically removed. Upon autocatalytic processing, the N-terminal threonine residues become exposed as the catalytically reactive residues, harboring both the nucleophile (the hydroxyl group) and the catalytic base (the N-terminal amine) involved in peptide bond cleavage. Each catalytic subunit has selectivity toward specific residues. β1 has caspase- or peptidyl-glutamyl peptidase-like activity, preferring cleavage at the C-terminus of acidic residues. β2 has trypsin-like activity and cleaves after basic residues, while β5 has chymotrypsin-like activity and prefers cleavage after hydrophobic residues. Figure \(5\) shows an interactive iCn3D model of the Human 20S Proteasome (6RGQ). The alpha subunits are shown in light gray and the beta subunits are in light cyan. Active site residue, as defined by databases, are shown for one of the alpha chain (L37, Q53, K55, H68, V170) and one of the beta chains (T1, D17, R19, K33, S130, D167, S170, G171). A possible generic mechanism for the cleavage of peptide bonds by the beta chain is shown in Figure \(6\). The mechanism shows how the newly exposed N-terminal threonine acts as both the nucleophile (the hydroxyl group) and the catalytic base. Proteins are normally targeted to the proteasome using ubiquitin labels attached covalently to a lysine residue, usually in a chained form that produces a polyubiquitinated protein. The process of protein polyubiquitination is carried out by a highly specialized and diverse enzymatic system, which includes families of ubiquitin-activating enzymes (E1), ubiquitin-conjugating enzymes (E2), and ubiquitin ligases (E3). The covalent attachment of ubiquitin to specific target proteins is mainly accomplished by stepwise enzymatic cascade reactions, and ubiquitin is attached to the substrates via the concerted action of ubiquitin-activating enzyme (E1), ubiquitin-conjugating enzyme (E2), and ubiquitin ligase (E3). The attachment of ubiquitin or ubiquitin chains to the substrate is a successive process, as shown in Figure \(7\). First, the C-terminal carboxylic acid is activated by adenylation using a molecule of ATP forming an adenylate (AMP-) intermediate. The adenylate acts as a good leaving group during the next reaction where an E1-ubiquitin thioester bond is formed between the C-terminal Gly carboxyl group of ubiquitin and the active site Cys of the E1 enzyme. AMP leaves the active site at this point. Note that ATP is used in many reactions to activate carboxylic acid functional groups through the formation of an adenylate intermediate and that this will be seen as a theme in many different types of reactions throughout this textbook. Once the ubiquitin is docked as a thioester on E1, it can be transferred to a Cys residue of the E2 enzyme to form an E2-ubiquitin thioester-linked intermediate. This enzymatic reaction is known as a transesterification. Eventually, the E2 transfers the ubiquitin to the substrate protein by E3. Ubiquitin is conjugated to the target protein through an isopeptide bond between its C-terminal glycine (Gly76) and the ε-amino group of a lysine residue. There are three typical ways of linking the ubiquitin with the substrate, as shown in Figure \(8\). The first is called mono-ubiquitination, which refers to the modification of one site of a substrate by a single ubiquitin molecule. The second is multi-mono-ubiquitination, which means adding several ubiquitin molecules repetitively to distinct sites (multi-mono-ubiquitination). The third is called polyubiquitination (including linear polyubiquitination and branched polyubiquitination), in which ubiquitin molecules are added to the same site (polyubiquitination, including linear polyubiquitination and branched polyubiquitination) of a substrate. In the second and third ways of linking, the previously attached ubiquitin serves as the “acceptor” of subsequently added ubiquitin. Of course, polyubiquitin chains linked by the same Lys are homogeneous, while those linked at different Lys are heterogeneous or mixed ones. The process of ubiquitination from activation to the attachment to the substrate is catalyzed by three major enzymes. The substrates labeled by ubiquitin are degraded by the 26S proteasome or play a non-degradative role in other processes. Abbreviations: APC, Anaphase-promoting complex; DUBs, Deubiquitinating enzymes; E1, Ubiquitin-activating enzyme; E2, Ubiquitin-conjugating enzyme; E3, Ubiquitin-ligase enzyme; Cul-based, Cullin-RING box1-Ligase; HECT, Homology to E6-AP C Terminus; Ub, Ubiqitin; SUMO, Small ubiquitin-related modifier; RBX1, RING-Box 1; RING, Really interesting new gene; RBR, RING1-IBR(cysteine/histidine-rich region)-RING2. Subsequently, the substrate complex tagged by the ubiquitin is either degraded by the 26S proteasome or executes nonproteolytic functions, such as the regulation of gene expression, cellular trafficking, or other biological function. In most cases, polyubiquitinated proteins are recognized and degraded by the 26S proteasome, and the ubiquitin or ubiquitin chain is hydrolyzed and freed by deubiquitinating enzymes (DUBs) for reuse in further conjugation cycles after being removed from the substrate protein (Figure 27.3.6). The family of E3 enzymes is large and diverse. It is estimated that there are 600-700 E3 enzymes in humans, representing approximately 5% of the human genome. Thus, E3 enzymes can be very substrate specific, leading to the specialized degradation of a small subset of proteins within the cell. E2 enzymes are the intersection between E1 and E3 enzymes and help to determine the ubiquitination of specific target proteins by interacting with different types of E3 enzymes. Figure \(9\) shows an interactive iCn3D model of the Yeast proteasome in resting state (C1-a) (6J2X). (long load) It's too complex to discuss this in much detail. The chains are presented in different colors. You should able to find the alpha and beta chains of the core 20S particle. The regulatory cap proteins comprising the lid are on top of the alpha ring.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/26%3A_Protein_Metabolism/26.03%3A_Translational_Regulation_and_Protein_Degradation.txt

Search Fundamentals of Biochemistry Introduction Each nucleated cell in a multicellular organism contains copies of the same DNA. Similarly, all cells in two pure bacterial cultures inoculated from the same starting colony contain the same DNA, except for changes that arise from spontaneous mutations. If each cell in a multicellular organism has the same DNA, then how is it that cells in different parts of the organism’s body exhibit different characteristics? Similarly, how is it that the same bacterial cells within two pure cultures exposed to different environmental conditions can exhibit different phenotypes? In both cases, each genetically identical cell does not turn on, or express, the same set of genes. Only a subset of proteins in a cell at a given time is expressed. Genomic DNA contains both structural genes, which encode products that serve as cellular structures or enzymes, and regulatory genes, which encode products that regulate gene expression. The expression of a gene is a highly regulated process. Whereas regulating gene expression in multicellular organisms allows for cellular differentiation, in single-celled organisms like prokaryotes, it primarily ensures that a cell’s resources are not wasted making proteins that the cell does not need at that time. Elucidating the mechanisms controlling gene expression is important to the understanding of human health. Malfunctions in this process in humans lead to the development of cancer and other diseases. Understanding the interaction between the gene expression of a pathogen and that of its human host is important for the understanding of a particular infectious disease. Gene regulation involves a complex web of interactions within a given cell among signals from the cell’s environment, signaling molecules within the cell, and the cell’s DNA. These interactions lead to the expression of some genes and the suppression of others, depending on circumstances. Prokaryotes and eukaryotes share some similarities in their mechanisms to regulate gene expression; however, gene expression in eukaryotes is more complicated because of the temporal and spatial separation between the processes of transcription and translation. Thus, although most regulation of gene expression occurs through transcriptional control in prokaryotes, regulation of gene expression in eukaryotes occurs at the transcriptional level and post-transcriptionally (after the primary transcript has been made). In bacteria and archaea, structural proteins with related functions are usually encoded together within the genome in a block called an operon and are transcribed together under the control of a single promoter, resulting in the formation of a polycistronic transcript, as shown in Figure \(1\). In this way, regulation of the transcription of all of the structural genes encoding the enzymes that catalyze the many steps in a single biochemical pathway can be controlled simultaneously, because they will either all be needed at the same time, or none will be needed. For example, in E. coli, all of the structural genes that encode enzymes needed to use lactose as an energy source are encoded next to each other in the lactose (or lac) operon under the control of a single promoter, the lac promoter. French scientists François Jacob (1920–2013) and Jacques Monod at the Pasteur Institute were the first to show the organization of bacterial genes into operons, through their studies on the lac operon of E. coli. For this work, they won the Nobel Prize in Physiology or Medicine in 1965. Each operon includes DNA sequences that influence its own transcription; these are located in a region called the regulatory region. The regulatory region includes the promoter and the region surrounding the promoter, to which transcription factors, proteins encoded by regulatory genes, can bind. Transcription factors influence the binding of RNA polymerase to the promoter and allow its progression to transcribe structural genes. A repressor is a transcription factor that suppresses the transcription of a gene in response to an external stimulus by binding to a DNA sequence within the regulatory region called the operator, which is located between the RNA polymerase binding site of the promoter and the transcriptional start site of the first structural gene. Repressor binding physically blocks RNA polymerase from transcribing structural genes. Conversely, an activator is a transcription factor that increases the transcription of a gene in response to an external stimulus by facilitating RNA polymerase binding to the promoter. An inducer, a third type of regulatory molecule, is a small molecule that either activates or represses transcription by interacting with a repressor or an activator. In prokaryotes, there are examples of operons whose gene products are required rather consistently and whose expression, therefore, is unregulated. Such operons are constitutively expressed, meaning they are transcribed and translated continuously to provide the cell with constant intermediate levels of the protein products. Such genes encode enzymes involved in housekeeping functions required for cellular maintenance, including DNA replication, repair, and expression, as well as enzymes involved in core metabolism. In contrast, other prokaryotic operons are expressed only when needed and are regulated by repressors, activators, and inducers. Prokaryotic operons are commonly controlled by the binding of repressors to operator regions, thereby preventing the transcription of the structural genes. Such operons are classified as either repressible operons or inducible operons. Repressible operons, like the tryptophan (trp) operon, typically contain genes encoding enzymes required for a biosynthetic pathway. As long as the product of the pathway, like tryptophan, continues to be required by the cell, a repressible operon will continue to be expressed. However, when the product of the biosynthetic pathway begins to accumulate in the cell, removing the need for the cell to continue to make more, the expression of the operon is repressed. Conversely, inducible operons, like the lac operon of E. coli, often contain genes encoding enzymes in a pathway involved in the metabolism of a specific substrate like lactose. These enzymes are only required when that substrate is available, thus expression of the operons is typically induced only in the presence of the substrate. The trp Operon - A Repressible Operon E. coli can synthesize tryptophan using enzymes that are encoded by five structural genes located next to each other in the trp operon, as shown in Figure \(2\). When environmental tryptophan is low, the operon is turned on. This means that transcription is initiated, the genes are expressed, and tryptophan is synthesized. However, if tryptophan is present in the environment, the trp operon is turned off. Transcription does not occur and tryptophan is not synthesized. When tryptophan is not present in the cell, the repressor by itself does not bind to the operator; therefore, the operon is active and tryptophan is synthesized. However, when tryptophan accumulates in the cell, two tryptophan molecules bind to the trp repressor molecule, which changes its shape, allowing it to bind to the trp operator. This binding of the active form of the trp repressor to the operator blocks RNA polymerase from transcribing the structural genes, stopping the expression of the operon. Thus, the actual product of the biosynthetic pathway controlled by the operon regulates the expression of the operon. The five structural genes needed to synthesize tryptophan in E. coli are located next to each other in the trp operon. When tryptophan is absent, the repressor protein does not bind to the operator, and the genes are transcribed. When tryptophan is plentiful, tryptophan binds the repressor protein at the operator sequence. This physically blocks the RNA polymerase from transcribing the tryptophan biosynthesis genes. Figure \(3\) shows an interactive iCn3D model of the E. Coli Trp repressor - operator complex (1TRO). The Trp repressor is shown as a dimer with one subunit gray and the other gold. The backbone of the two DNA strands is shown in spacefill magenta and cyan, except where the bases on the major grove interact with the Trp repressor. The tryptophan in each of the proton monomers is shown in spacefill with CPK colors and labeled. Noncovalent interactions (hydrogen bonds and salt bridges) between the protein and DNA are shown with dotted lines. 6 water-mediated hydrogen bonds to phosphate are not shown. Note that there are few H bonds between the protein and base hydrogen bond donors and acceptors, suggesting that the repressor might bind specifically through geometric interactions with the backbone along with the water-mediated hydrogen bonds. The Lac Operon: An Inducible Operon The lac operon is an example of an inducible operon that is also subject to activation in the absence of glucose. The lac operon encodes three structural genes, lacZ, lacY, and lacA, necessary to acquire and process the disaccharide lactose from the environment, as shown in Figure \(4\). Panel (A) shows a schematic representation of the lac operon in E. coli. The lac operon has three structural genes, lacZ, lacY, and lacA that encode for β-galactosidase, permease, and galactoside acetyltransferase, respectively. The promoter (p) and operator (o) sequences that control the expression of the operon are shown. Upstream of the lac operon is the lac repressor gene, lacI, controlled by the lacI promoter (p). Panel (B) shows the lac repressor inhibition of the lac operon gene expression in the absence of lactose. The lac repressor binds with the operator sequence of the operon and prevents the RNA polymerase enzyme which is bound to the promoter (p) from initiating transcription. Panel (C) shows that in the presence of lactose, some of the lactose is converted into allolactose, which binds and inhibits the activity of the lac repressor. The lac repressor-allolactose complex cannot bind with the operator region of the operon, freeing the RNA polymerase and causing the initiation of transcription. Expression of the lac operon genes enables the breakdown and utilization of lactose as a food source within the organism The lacZ gene encodes the β-galactosidase (β-gal) enzyme responsible for the hydrolysis of lactose into simple sugars glucose and galactose, as shown in Figure \(5\). The β-gal enzyme can also mediate the breakdown of the alternate substrate 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside (Xgal) (panel B). The breakdown product, 5-bromo-4-chloro-3-hydroxyindole – 1, spontaneously dimerizes to form the intensely blue product, 5,5′-dibromo-4,4′-dichloro-indigo – 2. Thus, Xgal has been a valuable research tool, not only in the study of the enzymatic activity of β-gal but also in the development of the commonly used blue-white DNA cloning system that utilizes the β-gal enzyme as a marker in molecular cloning experiments. The lac operon contains two more genes, in addition to lacZ (Fig. 4). The lacY gene encodes a permease that increases the uptake of lactose into the cell and lacA encodes a galactoside acetyltransferase (GAT) enzyme. The exact function of GAT during lactose metabolism has not been conclusively elucidated but acetylation is thought to play a role in the transport of the modified sugars. For the lac operon to be expressed, lactose must be present. This makes sense for the cell because it would be energetically wasteful to create the enzymes to process lactose if lactose was not available. In the absence of lactose, the lacI gene is constitutively expressed, expressing the lac repressor protein (Fig. 28.2.3 B). The lac repressor binds with an operator region of the lac operon and physically prevents RNA polymerase from transcribing the structural genes (Fig. 28.2.3 B). However, when lactose is present, the lactose inside the cell is converted to allolactose. Allolactose serves as an inducer molecule, binding to the repressor and changing its shape so that it is no longer able to bind to the operator DNA (Fig. 28.2.3 C). Removal of the repressor in the presence of lactose allows RNA polymerase to move through the operator region and begin transcription of the lac structural genes. In addition to lactose, laboratory experiments have revealed that the non-natural compound Isopropyl β-D-1-thiogalactopyranoside (IPTG) can also bind with the lac repressor and cause the expression of lac operon (panel C). Similar to Xgal, this compound has also been used as a research tool for molecular cloning. Figure \(6\) shows an interactive iCn3D model of the lactose operon repressor and its complexes with DNA (1LBG). The resolution of the structure above was insufficient to show the amino acid side chains. Figure \(7\) shows an interactive iCn3D model of the NMR solution structure of the dimer of LAC repressor DNA-binding domain complexed to its natural operator O1 (2KEI). Note the presents of white spheres representing hydrogen atoms (these don't appear in crystal structure but do in NMR structures. Color coding is the same as above. Zoom in to see specific interactions between the protein and the exposed DNA base hydrogen bond donors and acceptors. The complex of O1 and O2 shows similar specific and nonspecific contacts, which makes sense given the lambda repressor has similar affinity for those two operator sites. In contrast, one side of the O3 complex shows a loss of protein: DNA interactions, consistent with its lower affinity of its operator O3. The Lac Operon: Activation by Catabolite Activator Protein Bacteria typically can use a variety of substrates as carbon sources. However, because glucose is usually preferable to other substrates, bacteria have mechanisms to ensure that alternative substrates are only used when glucose has been depleted. Additionally, bacteria have mechanisms to ensure that the genes encoding enzymes for using alternative substrates are expressed only when the alternative substrate is available. In the 1940s, Jacques Monod was the first to demonstrate a preference for certain substrates over others through his studies of E. coli’s growth when cultured in the presence of two different substrates simultaneously. Such studies generated diauxic growth curves, like the one shown in Figure \(8\). Although the preferred substrate glucose is used first, E. coli grows quickly and the enzymes for lactose metabolism are absent. However, once glucose levels are depleted, growth rates slow, inducing the expression of the enzymes needed for the metabolism of the second substrate, lactose. Notice how the growth rate in lactose is slower, as indicated by the lower steepness of the growth curve. The ability to switch from glucose use to another substrate like lactose is a consequence of the activity of an enzyme called Enzyme IIA (EIIA). When glucose levels drop, cells produce less ATP from catabolism and EIIA becomes phosphorylated. Phosphorylated EIIA activates adenylyl cyclase, an enzyme that converts some of the remaining ATP to cyclic AMP (cAMP), a cyclic derivative of AMP and an important signaling molecule involved in glucose and energy metabolism in E. coli, as shown in Figure \(9\). As a result, cAMP levels begin to rise in the cell. This is an indicator to the cell, that overall energy levels are low and that ATP is being depleted. The lac operon also plays a role in this switch from using glucose to using lactose. When glucose is scarce, the accumulating cAMP caused by increased adenylyl cyclase activity binds to catabolite activator protein (CAP), also known as cAMP receptor protein (CRP). The complex binds to the promoter region of the lac operon, as shown in Figure \(10\). In the regulatory regions of these operons, a CAP binding site is located upstream of the RNA polymerase binding site in the promoter. The binding of the CAP-cAMP complex to this site increases the binding ability of RNA polymerase to the promoter region to initiate the transcription of the structural genes. Thus, in the case of the lac operon, for transcription to occur, lactose must be present (removing the lac repressor protein) and glucose levels must be depleted (allowing the binding of an activating protein). When glucose levels are high, there is catabolite repression of operons encoding enzymes for the metabolism of alternative substrates. Because of low cAMP levels under these conditions, there is an insufficient amount of the CAP-cAMP complex to activate the transcription of these operons. Figure \(11\) shows an interactive iCn3D model of the Catabolite activator protein CAP-DNA complex with bound cAMP (2CGP). Global Responses of Prokaryotes In prokaryotes, several higher levels of gene regulation have the ability to control the transcription of many related operons simultaneously in response to an environmental signal. A group of operons all controlled simultaneously is called a regulon. Alarmones When sensing impending stress, prokaryotes alter the expression of a wide variety of operons to respond in coordination. They do this through the production of alarmones, which are small intracellular nucleotide derivatives, such as guanosine pentaphosphate (pppGpp), as shown in Figure \(12\). Alarmones change which genes are expressed and stimulate the expression of specific stress-response genes. For example, pppGpp signaling is involved in the stringent response in bacteria, causing the inhibition of RNA synthesis when there is a shortage of amino acids present. This causes translation to decrease and the amino acids present are therefore conserved. Furthermore, pppGpp causes the up-regulation of many other genes involved in stress response such as the genes for amino acid uptake (from surrounding media) and biosynthesis. The use of alarmones to alter gene expression in response to stress appears to be important in pathogenic bacteria, as well. On encountering host defense mechanisms and other harsh conditions during infection, many operons encoding virulence genes are upregulated in response to alarmone signaling. Knowledge of these responses is key to being able to fully understand the infection process of many pathogens and to the development of therapies to counter this process. Quorum Sensing Quorum sensing (QS) is an intercellular communication mechanism of bacteria used to coordinate the activities of individual cells at the population level in response to surroundings through the production and perception of diffusible signal molecules such as Acyl Homoserine Lactones or small signaling peptides, as shown in Figure \(13\). The signal synthase, signal receptor, and signal molecules are three essential elements of the basic QS circuit machinery. Genes encoding signal-generating proteins are also included among the QS target genes. This forms an autoinduction feedback loop to modulate the generation of signal molecules. Several bacterial behaviors including virulence factors expression, secondary metabolites production, biofilm formation, motility, and luminescence are regulated by QS. Through complex regulatory networks, bacteria are capable of expressing corresponding genes according to their population size and of behaving in a coordinated manner. The left panel shows the typical Gram-negative quorum sensing mechanism. Acyl homoserine lactone molecules, synthesized by LuxI, passively pass the bacterial cell membrane and when a sufficient concentration is reached (threshold level) activate the intracellular LuxR which subsequently activates target gene expression in a coordinated way. Note that a single cell is shown for simplicity. However, acyl homoserine lactones will commonly diffuse and target neighboring cells within the colony to mediate a communal or population response within the bacterial colony. The right panel shows that quorum-sensing peptides are synthesized by the bacterial ribosomes as pro-peptidic proteins and undergo posttranslational modifications during excretion by active transport. The quorum-sensing peptides bind membrane-associated receptors which get autophosphorylated and activate intracellular response regulators via phosphotransfer. These phosphorylated response regulators induce increased target gene expression. For example, some microbial species, such as Staphylococcus aureus, can encase their community within a self-produced matrix of hydrated extracellular polymeric substances that include polysaccharides, proteins, nucleic acids, and lipid molecules. These encasements are known as biofilms. The formation of the biofilm on solid surfaces is a step-wise process comprising several stages, as shown in Figure \(14\). It starts with the conditioning of the surface through the coating with macromolecules from the aqueous surrounding, which enables the initial reversible adhesion of microorganisms. The next step is the formation of stronger, irreversible attachments to the surface, followed by the proliferation and aggregation of microorganisms into multicellular and multilayered clusters, which actively produce an extracellular matrix. Some cells in the mature biofilms continuously detach and separate from the aggregates, representing a continuous source of planktonic bacteria that can subsequently spread and form new microcolonies. Biofilms are a common cause of chronic, nosocomial (originating in a hospital), and medical device-related infections because they can develop either on vital or necrotic tissue as well as on the inert surfaces of different implanted materials. Moreover, biofilms are linked with high-level resistance to antimicrobials, frequent treatment failures, and increased morbidity and mortality. As a consequence, biofilm infections and accompanying diseases have become a major health concern and a serious challenge for both modern medicine and pharmacy. The rough estimation shows that more than 60% of hospital-associated infections are attributable to the biofilms formed on indwelling medical devices, which result in more than one million cases of infected patients annually and more than \$1 billion in hospitalization costs per year in the USA. Biofilm infections share some common characteristics: slow development in one or more hot spots, delayed clinical manifestation, and persistency for months or years, usually with interchanging periods of acute exacerbations and absence of clinical symptoms. Even though they are less aggressive than acute infections, their treatment is challenging to a greater extent. There is upto a 1000-fold decrease in the susceptibility of biofilms to antimicrobial agents and disinfectants as well as resistance to host immune response. Thus, ways to reduce or inhibit biofilm formation are highly sought. The majority of the proposed biofilm-control methods focus on: (i) prevention and minimization of biofilm formation by selection and surface modifications of anti-adhesive materials; (ii) debridement techniques including ultrasound and surgical procedures; (iii) disruption of biofilm QS-signaling system; or (iv) achieving proper drug penetration and delivery to formed biofilms by the use of an electromagnetic field, ultrasound waves, photodynamic activation or specific drug delivery systems. Alternate σ Factors Since the σ subunit of bacterial RNA polymerase confers specificity as to which promoters should be transcribed, altering the σ factor used is another way for bacteria to quickly and globally change what regulons are transcribed at a given time. The σ factor recognizes sequences within a bacterial promoter, so different σ factors will each recognize slightly different promoter sequences. In this way, when the cell senses specific environmental conditions, it may respond by changing which σ factor it expresses, degrading the old one and producing a new one to transcribe the operons encoding genes whose products will be useful under the new environmental condition. For example, in sporulating bacteria of the genera Bacillus and Clostridium (which include many pathogens), a group of σ factors controls the expression of the many genes needed for sporulation in response to sporulation-stimulating signals. Prokaryotic Attenuation and Riboswitches Although most gene expression is regulated at the level of transcription initiation in prokaryotes, there are also mechanisms to control both the completion of transcription, as well as translation, concurrently. Since their discovery, these mechanisms have been shown to control the completion of transcription and translation of many prokaryotic operons. Because these mechanisms link the regulation of transcription and translation directly, they are specific to prokaryotes, because these processes are physically separated in eukaryotes. One such regulatory system is attenuation, whereby secondary stem-loop structures formed within the 5’ end of an mRNA being transcribed determine if transcription to complete the synthesis of this mRNA will occur and if this mRNA will be used for translation. Beyond the transcriptional repression mechanism already discussed, attenuation also controls the expression of the trp operon in E. coli as shown in Figure \(15\). The trp operon regulatory region contains a leader sequence called trpL between the operator and the first structural gene, which has four stretches of RNA that can base pair with each other in different combinations. When a terminator stem-loop forms, transcription terminates, releasing RNA polymerase from the mRNA. However, when an antiterminator stem-loop forms, this prevents the formation of the terminator stem-loop, so RNA polymerase can transcribe the structural genes. When tryptophan is plentiful, translation of the short leader peptide encoded by trpL proceeds, the terminator loop between regions 3 and 4 forms, and transcription terminates. When tryptophan levels are depleted, translation of the short leader peptide stalls at region 1, allowing regions 2 and 3 to form an antiterminator loop, and RNA polymerase can transcribe the structural genes of the trp operon. A related mechanism of concurrent regulation of transcription and translation in prokaryotes is the use of a riboswitch, a small region of noncoding RNA found within the 5’ end of some prokaryotic mRNA molecules, as shown in Figure \(16\). A riboswitch may bind to a small intracellular molecule to stabilize certain secondary structures of the mRNA molecule. The binding of the small molecule determines which stem-loop structure forms, thus influencing the completion of mRNA synthesis and protein synthesis. Riboswitches found within prokaryotic mRNA molecules can bind to small intracellular molecules, stabilizing certain RNA structures, and influencing either the completion of the synthesis of the mRNA molecule itself (left) or the protein made using that mRNA (right). Figure \(17\) shows interactive iCn3D models of a series of bacterial riboswitches. They are described in the legend below. Guanine-responsive riboswitch bound to metabolite hypoxanthine (4FE5) A. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...Nh1Sv7oSSyC6Z6 Divalent cation-sensing regulatory RNA (2QBZ) B. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...VbU9Vf6DUA4VE8 Cyclid-di-GMP RNA riboswitch (3IRW) C. (Copyright; author via source). Click the image for a popup or use this external link:https://structure.ncbi.nlm.nih.gov/i...wVntbdScP3VCF8 GlmS ribozyme bound to glucosamine-6-phosphate (2Z75) D. (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...w9T4K9ffK1NU27 A: Guanine-responsive riboswitch bound to metabolite hypoxanthine (4FE5) - Hypothanine, involved in purine metabolism, is shown bound to RNA representing the 5' untranslated region of the xanthine phosphoribosyltransferase (xbt)/ xanthine-specific purine permease (pbux) genes that lead to transcription termination. B: The M-box in mycobacterial genes regulating Mg2+ transport binds divalent cation. They are transcribed under low Mg2+ concentrations. Salt bridges (ion-ion interactions) are shown in cyan and pi-cation interactions in red dotted lines C: Bis-(3'-5')-cyclic dimeric guanosine monophosphate (c-di-GMP) is a second messenger in bacteria and regulates many cellular processes including the formation of biofilms. The riboswitch shown here is from Vibrio cholerae. The U1 small nuclear ribonucleoprotein A is shown in cyan. Figure \(18\) shows a cartoon of the actual riboswitch in the 5' untranslated region of target genes D. This ribozyme is in the 5′ untranslated region of glucosamine-6-phosphate synthase mRNA. The protein enzyme, 2. This protein enzyme catalyzes the conversion of fructose 6-phosphate and glutamine to glucosamine 6-phosphate (GlcN6P) and glutamate. The glmS ribozyme in the 5'-untranslated region cleaves itself on binding GlcN6P. This self-cleavage is inhibited by glucose 6-phosphate (Glc6P). Hence high levels of the gene product for the synthase lead to cleavage of its own mRNA. The glmS ribozyme RNA is shown in gray and its substrate RNA in cyan.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/27%3A_Regulation_of_Gene_Expression/27.01%3A_Regulation_of_Gene_Expression_in_Bacteria.txt

Search Fundamentals of Biochemistry As seen in Chapter 26, the initiation of transcription requires the assembly of a multitude of transcription factors (TF) localized at the promoter region. Transcription can also utilize far-reaching interactions of enhancers, that bind at a distant DNA site and loop back around to stabilize the RNA polymerase at the promoter. Control of transcriptional initiation is dependent on TF factor activation, TF binding with specific DNA recognition sequences, and chromatin remodeling. Transcription Factor (TF) Activation Many TFs are expressed within cells and held in an inactive conformation until the right environmental stimulus is present within the cell. Cellular signaling pathways can cause post-translational protein modifications leading to TF activation or small molecules may physically bind and allosterically modify the protein structure to mediate activation. Here we will use examples from the cell cycle signaling cascade and steroid hormone receptor pathways to highlight some mechanisms of TF activation. A key element to take away from this section is that transcription factor activation is often highly pleiotropic and has many cellular effects. Depending on the cell type and the environmental conditions, different combinations of downstream target genes may be activated or inactivated. Teasing apart these intricacies and the physiological effects that they have within an organism is a major goal of ongoing research. Cell Cycle Regulation by p53 p53 is one of the most studied proteins in science. To date, over 68,000 papers appear in PubMed containing p53 or TP53 in the title and/or abstract. Originally described as an oncogene (since a mutated, functionally altered form of the protein was first characterized), p53 is now recognized as the most frequently inactivated tumor suppressor in human cancers. It is a transcription factor that controls the expression of genes and miRNAs affecting many important cellular processes including proliferation, DNA repair, programmed cell death (apoptosis), autophagy, metabolism, and cell migration, as shown in Figure \(1\). Many of those processes are critical to a variety of human pathologies and conditions extending beyond cancer, including ischemia, neurodegenerative diseases, stem cell renewal, aging, and fertility. Notably, p53 also has non-transcriptional functions, ranging from intrinsic nuclease activity to activation of mitochondrial Bak (Bcl-2 homologous antagonist killer) and caspase-independent apoptosis. As a transcription factor, p53 responds to various genotoxic insults and cellular stresses (e.g., DNA damage or oncogene activation) by inducing or repressing the expression of over a hundred different genes. p53 transcriptional regulation plays a dominant role in causing the arrest of damaged cells, facilitating their repair and survival, or inducing cell death when DNA is damaged irreparably. p53 can also cause cells to become permanently growth arrested, and there is compelling in vivo evidence that these “senescent” cells secrete factors that enhance their clearance by the immune system, leading to tumor regression. Through these mechanisms, p53 helps maintain genomic stability within an organism, justifying its long-held nickname “guardian of the genome”. Other p53 gene targets are involved in inhibiting tumor cell angiogenesis, migration, metastasis, and other important processes (such as metabolic reprogramming) that normally promote tumor formation and progression Figure \(2\) shows an interactive iCn3D model of the human p53 tetramer bound to the natural CDKN1A(p21) p53-response element (3TS8). Each monomer of the p53 tetramer is shown in a different color. The noncovalent interactions of the brown monomer with the DNA are shown, with key amino acids and nucleic acid bases shown in CPK-colored sticks.  They interact with the Zn2+ ions, shown in the light green monomer. Normally, p53 levels are kept low by its major antagonist, Mdm2, an E3 ubiquitin ligase that is itself a transcriptional target of p53. Stress signals, such as DNA damage, oncogene activation, and hypoxia, promote p53 stability and activity by inducing post-translational modifications (PTMs) and tetramerization of p53. p53 functions as a transcription factor that binds to specific p53 response elements upstream of its target genes. p53 affects many important cellular processes linked to tumor suppression, including the induction (green) of senescence, apoptosis, and DNA repair as well as inhibition (red) of metabolism, angiogenesis, and cell migration. These functions are largely mediated through transcriptional regulation of its targets (examples given). p53 protein function is regulated post-translationally by coordinated interaction with signaling proteins including protein kinases, acetyltransferases, methyl-transferases, and ubiquitin-like modifying enzymes as shown in Figure \(3\). The majority of the sites of covalent modification occur at intrinsically unstructured linear peptide docking motifs that flank the DNA-binding domain of p53 which plays a role in anchoring or in allosterically activating the enzymes that mediate covalent modification of p53. In undamaged cells, the p53 protein has a relatively short half-life and is degraded by a ubiquitin-proteasome-dependent pathway through the action of E3 ubiquitin ligases, such as MDM2 (Fig 28.3.1). Following stress, p53 is phosphorylated at multiple residues, thereby modifying its biochemical functions required for increased activity as a transcription factor. Post-translational modifications help to stabilize the tetramer formation of the protein and enhance the translocation of the protein from the cytoplasm into the nucleus. The tetrameric form of p53 is then functional to bind to DNA in a sequence-specific manner and either activate or repress transcription, depending on the target sequence. Some post-translational modifications, such as acetylation, are DNA-dependent and can play a role in chromatin remodeling and activation of p53 target gene expression. It should be noted that single-point mutations that modify the ability of the protein to be phosphorylated in one position, typically do not show a decrease in the stabilization or activation of the protein following a damage or stress event. Thus, multiple modifications likely allow for redundancy within this pathway and ensure the activation of the protein following a stress event. Furthermore, the environment within the cell can lead to different p53 phenotypes, such as the activation of growth arrest and DNA repair processes (ie if there is not a lot of damage) or it can lead to the activation of apoptosis or programmed cell death pathways (ie if the damage is too extensive to be repaired). Steroid Hormone Receptors Steroid hormone receptors (SHRs) belong to the superfamily of nuclear receptors (NRs), which are one of the essential classes of transcriptional factors. NRs play a critical role in all aspects of human development, metabolism.  and physiology. Since they generally act as ligand-activated transcription factors, they are an essential component of cell signaling. NRs form an ancient and conserved family that arose early in the metazoan lineage. NR molecular evolution is characterized by major events of gene duplication and gene losses. Phylogenetic analysis revealed a distinct separation of NR ligand binding domains (LBDs) into 4 monophyletic branches, the steroid hormone receptor-like cluster, the thyroid hormone-like receptors cluster, the retinoid X-like and steroidogenic factor-like receptor cluster and the nerve growth factor-like/HNF4 receptor cluster, as shown in Figure \(4\). Here we will focus on the Steroid Hormone-Like Receptors branch (SHRs). SHRs plays a key role in many important physiological processes like organ development, metabolite homeostasis, and response to external stimuli. The estrogen receptor comes in two major forms, ERα and ERβ. Other members of this subgroup include the cortisol-binding glucocorticoid receptor (GR), the aldosterone-binding mineralocorticoid receptor (MR), the progesterone receptor (PR), and the dihydrotestosterone (DHT) binding androgen receptor (AR), as shown in Figure \(5\ below. Panel A shows a phylogenetic tree of the Steroid Hormone Receptor (SHR) family showing the evolutionary interrelationships and distance between the various receptors. Based on alignments available at The NucleaRDB [Horn et al., 2001]. Panel B shows that all steroid receptors are composed of a variable N-terminal domain (A/B) containing the AF-1 transactivation region, a highly conserved DNA Binding Domain (DBD), a flexible hinge region (D), and a C-terminal Ligand Binding Domain (LBD, E) containing the AF-2 transactivation region. The estrogen receptor α is unique in that it contains an additional C-terminal F domain. Numbers represent the length of the receptor in amino acids. The members of the Steroid Hormone Receptor family share a similar, modular architecture, consisting of several independent functional domains (Fig. 5B above). Most conserved is the centrally located DNA binding domain (DBD) containing the characteristic zinc-finger motifs. The DBD is followed by a flexible hinge region and a moderately conserved Ligand Binding Domain (LBD), located at the carboxy-terminal end of the receptor. The estrogen receptor α is unique in that it contains an additional F domain of which the exact function is unclear. The LBD is composed of twelve α-helices (H1-H12) that together fold into a canonical α-helical sandwich. Besides its ligand binding capability, the LBD also plays an important role in nuclear translocation, chaperone binding, receptor dimerization, and coregulator recruitment through its potent ligand-dependent transactivation domain, referred to as AF-2. A second, ligand-independent, transactivation domain is located in the more variable N-terminal part of the receptor, designated as AF-1. To date, no crystal structure of a full-length SHR exists, though structures of the DBD and LBD regions of most SHRs are available. These have helped significantly in understanding the molecular aspects of DNA and ligand binding, but have to some extent also led to biased attention to these parts of the receptor only. For example, many coregulator interaction studies are still performed with the LBD only, while numerous studies have demonstrated that the AF-2 domain often tells only part of the story. With the help of biophysical techniques, however, it is feasible to study the full-length receptor in its native environment. Most SHRs remain in the cytoplasm of the cell until they are bound with the appropriate steroid as shown in Figure \(6\). Steroid binding causes the dimerization of SHRs and localization to the cell nucleus, where the SHRs interact with the DNA at sequence-specific motifs known as Hormone Response Elements (HREs) (Step 5). Many SHRs can also interact with membrane-bound receptors and affect cellular signaling pathways, in addition to the activation of gene expression (step 6). Steroid hormones, such as estrogens, reach their target cells via the blood, where they are bound to carrier proteins. Naturally occurring estrogens including estradiol, estrone, estriol, differ primarily in structure on the presence of hydroxyl groups (Fig. 28.3.6). Estradiol is the predominant estrogen during reproductive years both in terms of absolute serum levels as well as in terms of estrogenic activity. During menopause, estrone is the predominant circulating estrogen, and during pregnancy, estriol is the predominant circulating estrogen in terms of serum levels. Another type of estrogen called estetrol (E4) is produced also produced predominantly during pregnancy as shown in Figure \(7\). Estrogens function in many physiological processes, including the regulation of the menstrual cycle and reproduction, maintaining bone density, brain function, cholesterol mobilization, and maturation of reproductive organs during development, and they play a role in controlling inflammation. Because of their lipophilic nature, it is thought that steroid hormones, such as estrogen, pass the cell membrane by simple diffusion, although some evidence exists that they can also be actively taken up by the endocytosis of carrier protein-bound hormones. For a long time, it has been assumed that binding of the ligand resulted in a simple on/off switch of the receptor (Fig. 6, step 1). While this is likely the case for typical agonists like estrogen and progesterone, this is not always correct for receptor antagonists, used in drug therapy. These antagonists come in two kinds, so-called partial antagonists (for the estrogen receptors known as SERMs for Selective Estrogen Receptor Modulators) and full antagonists. The partial antagonist can, depending on cell type, act as a SHR agonist or antagonist. In contrast, full antagonists (for ER known as SERDs for Selective Estrogen Receptor Downregulators) always inhibit the receptor, independent of cell type, in part by targeting the receptor for degradation. Binding of either type of antagonist results in major conformational changes within the LBD and in the release from heat shock proteins that thus far had protected the unliganded receptor from unfolding and aggregation (Fig. 6 step 2). Figure \(8\) shows an interactive iCn3D model of the Androgen Receptor DNA-Binding Domain Bound to a Direct Repeat Response Element (1R4I). Transcription Factor (TF) Recognition and Binding to DNA TF controls gene expression by binding to their target DNA site to recruit, or block, the transcription machinery onto the promoter region of the gene of interest. Their function relies on the ability to find their target site quickly and selectively. In living cells, TFs are present in nM concentrations and bind the target site with comparable affinity, but they also bind any DNA sequence (nonspecific binding), resulting in millions of low affinity (i.e., >10−6 M) competing sites. Nonspecific binding facilitates the search for the target site by three major mechanisms as shown in Figure \(9\). The second scenario is a ‘hopping’ mechanism, in which a TF might hop from one site to another in 3D space by dissociating from its original site and subsequently binding to the new site. This may happen within the same chain and re-association occurs adjacent to the former dissociated site. A third search mechanism is described as ‘intersegmental transfer’. In this scenario, the protein moves between two sites via an intermediate ‘loop’ formed by the DNA and subsequently binds at two different DNA sites. This mechanism applies to TFs with two DNA-binding sites. Proteins with two DNA-binding sites can occasionally bind non-specifically to two locations situated far apart within the DNA strand, that are brought into close contact through the formation of these loops. Such TFs transfer across a point of close contact without dissociating from the DNA. Top: When the transcription factor (pink ring) moves from one site to another by sliding along the DNA and is transferred from one base pair to another without dissociating from the DNA, this mechanism is called sliding. Center: Hopping occurs when the transcription factor moves on the DNA by dissociating from one site and re-associating with another site. Bottom: Intersegmental transfer describes the mechanism by which the transcription factor gets transferred through DNA bending or the formation of a DNA loop, resulting in the protein being bound transiently to both sides and subsequently moving from one site to the other. One of the main scenarios involves a ‘sliding’ mechanism, in which the protein moves from its initial non-specific site to its actual target site by sliding along the DNA (also known as 1-dimensional (1D) sliding). When the TF starts to move and shift counterions from the phosphate backbone, the same number of counterions binds to the site left free by the protein. The sliding rate is also dependent on the hydrodynamic radius of the protein; the required rotational movement over the DNA backbone is greater for larger proteins, that tend to slide slowly. Recent Updates:  9/25/23 The sliding model that was proposed by Von Hippel and Berg suggests that DNA-binding protein exists in two interconverting conformations.  One is a specific form (O) that can bind to a target DNA sequence, such as an operator in DNA, through specific hydrogen bonds (along with electrostatic interaction) characterized by a low KD.  The other is a nonspecific form (D) that binds mainly with weaker affinity through electrostatic interactions and a high KD. Nonspecific binding brings the protein to the DNA surface.  Dynamic conformational changes from the O to D allow sampling of hydrogen bonds between donor and acceptors in the protein and in the major grove of the protein.  The protein can diffuse much more quickly along the DNA to find its target site since the search for the specific target site is now effectively 1D instead of 3D.  There is no thermodynamic barrier to sliding since counterions that leave the DNA when bound to the protein rebind behind it as the protein slides.  These processes are illustrated in Figure \(10\) below. Figure \(10\): Two-state model sliding model of DNA protein binding to its specific binding site (operator) There are a large number of overlapping nonspecific binding sites (let's say each is 6 base pairs in length), which also help drive the nonspecific binding of the protein to the DNA through entropy increases (i.e consider the probability of binding to 1 site on the DNA versus multitudes of overlapping sites).  Experiments show that the on-rate for a DNA binding protein for finding its target site increases with increasing length of the DNA molecule the specific site is embedded in.  The opposite would be expected given the diffusion rate of large molecules.  In fact, the kon was found to be greater than diffusion-controlled limits, which can be explained by the reduced dimensionality of the search for the specific site when the protein is loosely bound through nonspecific electrostatic interactions that enable the 1D search. Each eukaryotic TF controls tens to hundreds of genes scattered throughout the genome, and expressing each gene needs various TFs simultaneously binding to their sites to form the transcription complex, an extremely rare event in probabilistic terms. As a result, the in vivo site occupancy patterns of eukaryotic TFs are more complex than predicted by their in vitro site-specific binding profiles and do not strongly correlate with the actual levels of gene expression. An interesting feature highlighted by genome analysis is an accumulation of potential TF binding sites in regions flanking eukaryotic genes. Such clusters of degenerate recognition sites are assumed to be key for transcription control and thus are generally classified as gene regulatory regions (RR). For example, the affinity of the Drosophila TF Engrailed to the RRs of its target genes is strongly amplified by long tracts of degenerate consensus repeats that are present in such regions. Role of Short Tandem Repeats (STRs) in the Genome As we have mentioned previously, only about 1.5% of the human genome encodes genes for actual proteins.  Much of the genome is transcribed at low levels into RNAs, some of which have clearly defined functions (examples include rRNA, mRNAs, and regulatory RNAs).  A large part is presumably involved in facilitating the 3D organization of the genome and its dynamic architecture, which determines its replicative and transcriptive access. One poorly understood feature of genomic DNA is short tandem repeats (STRs).  These repeats stretch up to 100 nucleotides in length with each repetitive tandem repeat running from just 1 to 6 bases long.  They comprise about 6% of the genome (compared to the 1.5% for protein-coding genes), and are found in abundance in chromatin that is transcribed mRNA for proteins. For example, a specific DNA sequence might be GTCACGTGAC while a small STR would be (CG)6C(CG)11 Specifically, STRs surround sites where classic transcription factors (TF) bind.  As we described in this and the previous chapter sections, TFs bind through specific DNA binding motifs (like helix-loop-helix or Zn2+ fingers) to consensus sequences (such as response elements and enhancers) in the DNA as they function to control transcription.  The binding of transcription factors or other proteins to target sequences occurs initially through nonspecific electrostatic interactions, followed by a dimensionally restricted diffusion along the DNA as the protein finds its specific sequence target. In contrast, the STRs offer little sequence uniqueness for high affinity, low KD binding sites for specific protein interactions, so the question remains as to how they express function, which their omnipresence suggests they have.  Studies by Horton et al (Science 381, 1304 (2023) have shown that classic TFs do indeed bind STRs, albeit at lower affinity (higher KD) compared to their binding to classic TF-specific sequences.  The bind with higher affinity than to nonspecific DNA sequences.  Just as nonspecific interactions facilitate TF binding to promoter sites, so do the multiple STRs that straddle the DNA binding element. If a TF binds its isolated target DNA with a low effective KD (high affinity) and -ΔG0 value, a target DNA surrounded by multiple STRs would have an even lower effective KD (higher affinity) and even more -ΔG0 value.  They do so by increasing the effective on rate (kon) for protein binding.  (Remember that KD = koff/kon. )  The effective size of the target for the TF becomes greater when it is an “island” in the middle of a STR “sea”. The increased affinity stems in large part from the more favorable entropy of having the TF bind not to just 1 site but effectively to multiple overlapping sites.  This also increases the localized concentration of TF near the specific site which drives binding.  The koff is not expected to change. Another effect of the STRs on TF binding is that multiple DNA-binding proteins can bind to the same site through their interactions with STRs at the site, leading to new ways to regulate gene transcription.  The group studies just two TFS but sequence analyses suggest that many TF would use a similar mechanism. Histone Modification and Chromatin Remodeling Regulation of transcription involves dynamic rearrangements of chromatin structure. Recall that eukaryotic DNA is complexed with histone octamers, which are composed of dimers of the core histones H2A, H2B, H3, and H4. 147 bp of DNA are wrapped 1.65 times around each octamer forming nucleosomes, the basic packaging units of chromatin. Nucleosomes, connected by linker DNA of variable length as “beads on a string”, generate the 11 nm linear structure. The linker histone H1 is positioned at the top of the core histone octamer and enables higher organized compaction of DNA into transcriptionally inactive 30 nm fibers. To understand the role of chromatin in the regulation of transcription it is important to know where nucleosomes are positioned and how the positioning is achieved. Basically, there are four groups of activities that change chromatin structure during transcription: (1) histone modifications, (2) eviction and repositioning of histones, (3) chromatin remodeling, and (4) histone variant exchange. Histone modifiers introduce post-translational, covalent modifications to histone tails and thereby change the contact between DNA and histones. These modifications govern access to regulatory factors. Histone chaperones aid in the eviction and positioning of histones. A third class of chromatin restructuring factors is ATP-dependent chromatin remodelers. These multi-subunit complexes utilize energy from ATP hydrolysis for various chromatin remodeling activities including nucleosome sliding, nucleosome displacement, and the incorporation and exchange of histone variants. Post-translational modifications (PTMs) of histone proteins are a primary mechanism that controls chromatin architecture. Over 20 distinct types of histone PTMs have been described, among which the most abundant ones are acetylation and methylation of lysine residues. Histone PTMs can be deposited on and removed from chromatin by different enzymes, known as histone PTM ‘writers’ and ‘erasers’. Histone PTMs exert their regulatory effects via two main mechanisms. First, histone PTMs serve as docking sites for various nuclear proteins––histone PTM ‘readers’––that specifically recognize modified histone residues through their modification-binding domains. Recruitment of these proteins at specific genomic loci promotes key chromatin processes, such as transcriptional regulation and DNA damage repair. Second, some histone PTMs, such as acetylation, directly affect chromatin's higher-order structure and compaction, thereby controlling chromatin accessibility to protein machinery such as those involved in transcription. Chromatin may adopt one of two major states interchangeably. These states are heterochromatin and euchromatin. Heterochromatin is a compact form that is resistant to the binding of various proteins, such as transcriptional machinery. In contrast, euchromatin is a relaxed form of chromatin that is open to modifications and transcriptional processes, as shown in Figure \(10\). Histone methylation promotes the formation of Heterochromatin whereas, histone acetylation promotes euchromatin. The addition of methyl groups to the tails of histone core proteins leads to histone methylation, which in turn leads to the adoption of a condensed state of chromatin called ‘heterochromatin.’ Heterochromatin blocks transcription machinery from binding to DNA and results in transcriptional repression. The addition of acetyl groups to lysine residues in the N-terminal tails of histones causes histone acetylation, which leads to the adoption of a relaxed state of chromatin called ‘euchromatin.’ In this state, transcription factors and other proteins can bind to their DNA binding sites and proceed with active transcription. Chromatin remodeling can also be an ATP-dependent process and involve histone dimer ejection, full nucleosome ejection, nucleosome sliding, and histone variant exchange as shown in Figure \(12\). ATP-dependent chromatin remodeling complexes bind to nucleosome cores and the surrounding DNA, and, using energy from ATP hydrolysis, they disrupt the DNA-histone interactions, slide or eject nucleosomes, alter nucleosome structures, and modulate the access of transcription factors to the DNA (Figure 28.3.9). In addition to modulating gene expression, some of the complexes are involved in nucleosome assembly and organization, following transcription at locations in which nucleosomes have been ejected, packing of DNA, following replication, and DNA repair. Panel (a) shows a subset of ISWI and CHD complexes is involved in nucleosome assembly, maturation, and spacing. Panel (b) shows SWI/SNF complexes are primarily involved in histone dimer ejection, nucleosome ejection, and nucleosome repositioning through sliding, thus modulating chromatin access. Panel (c) shows INO80 complexes are involved in histone exchange. It should be noted that the complexes might be involved in other chromatin remodeling functions. Figure \(13\)s shows the effects of Histone Variant H3.3 on C. elegans Lifespan Protein-DNA Interactions Proteins use a wide range of DNA-binding structural motifs, such as homeodomain (HD), helix-turn-helix (HTH), and high-mobility group box (HMG) to recognize DNA. HTH is the most common binding motif and can be found in several repressor and activator proteins, as shown in Figure \(14\). Despite their structural diversity, these domains participate in a variety of functions that include acting as substrate interaction mediators, enzymes to operate DNA, and transcriptional regulators. Several proteins also contain flexible segments outside the DNA-binding domain to facilitate specific and non-specific interactions. For example, many HD proteins use N-terminal arms and a linker region to interact with DNA. The Encyclopedia of DNA Elements (ENCODE) data suggest that about 99.8% of putative binding motifs of TFs are not bound by their respective TFs in the genome. It is, therefore, clear that the presence of a single binding motif per TF is not adequate for TF binding. Figure \(14\) shows interactive iCn3D models of the transcription factor binding domains as depicted in the figure above. (Copyright; author via source) POU protein:DNA complex HTH-HD domain (3l1p) Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...fHmWMvUCiJus77 Human Hsf1 with Satellite III repeat DNA - HTH Domain (5d5v) Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...qkHMwpH7LdDmr7 POU-HMG-DNA ternary complex - HTM-HMG domain (1gt0) Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...b9X7DEGSK2n2V7 Klf4 zinc finger DNA binding domain in complex with methylated DNA(4m9e) Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...T7ZXxMhQ66WNL9 Lactose operon repressor and its complexes with DNA and inducer (1lbg) Click the image for a popup or use this external link:https://structure.ncbi.nlm.nih.gov/i...6v3y24U7N2qYC8 Myc-Max and Mad-Max recognizing DNA(1nkp) Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...qcTBN4UwxPFH99 Most of the search mechanistic studies that try to determine how TFs find their binding sites are limited to naked DNA-protein complexes, which do not reflect the actual crowded environment of a cell. Studies with naked DNA and transcription factors have shown that many DNA-binding proteins travel a long distance by 1D diffusion. However, the search process for eukaryotes must occur in the presence of chromatin, which can hinder protein mobility. In this case, the protein must dissociate from the DNA, enter a 3D mode of diffusion state, and continue the target site searching process. The sliding and intersegmental transfer mechanisms can be explained through the example of the lac repressor. The lac repressor contains 4 identical monomers (a dimer of dimers) for its DNA binding. The binding sequence of these dimers is symmetric or pseudo-symmetric, and each half is identified by these identical monomers. The HTH domain of the lac repressor is the DNA-binding domain that facilitates the interaction with its target site on DNA as shown in Figure \(15\). As a result of a rapid search (sliding) along the DNA molecule and intersegmental transfer between distant DNA sequences, the lactose repressor finds its target sites faster than the diffusion limit. The section comprised between residues 1–46 of the HTH protein domain, characterized by three α-helices, maintains its secondary structure through specific and non-specific binding. When the repressor binds to a non-specific site, the HTH domain interacts with the DNA backbone and maintains the interaction with its helix region in the major groove juxtaposition. This arrangement facilitates the interaction of the recognition helix with the edges of the DNA bases, enabling the repressor to walk or search for its specific site on the DNA. The C-terminal residues of the DNA-binding domain, residues 47–62, form the hinge region, and are normally disordered during non-specific recognition; however, during specific site recognition, residues 50–58 acquire an α-helix configuration (hinge helix) (Fig. 15 above). The disordered hinge region and the flexibility of the HTH domain allow the protein to move freely along the DNA to search for its target site. In specific binding complexes, the hinge helix of each monomer is located at the symmetrical center of the binding site, thereby causing the hinge helices to interact with each other (intersegmental transfer) to allow better stability. Moreover, DNA bends at the symmetrical center of the specific binding site (37° angle), thereby supporting monomer-monomer interactions. In addition to the helix-turn-helix structure, the zinc finger motif is also very common, especially in eukaryotic TFs, as shown in Figure \(16\). Proteins that contain zinc fingers (zinc finger proteins) are classified into several different structural families. Unlike many other clearly defined supersecondary structures such as Greek keys or β hairpins, there are a number of types of zinc fingers, each with a unique three-dimensional architecture. A particular zinc finger protein’s class is determined by this three-dimensional structure, but it can also be recognized based on the primary structure of the protein or the identity of the ligands coordinating the zinc ion. Despite the large variety of these proteins, however, the vast majority typically function as interaction modules that bind DNA, RNA, proteins, or other small, useful molecules.  Variations in structures serve primarily to alter the binding specificity of a particular protein. The most common type of zinc finger motif utilizes two Cys and two His residues (CCHH) coordinating the Zn(II) ion to adopt a ββα fold with three hydrophobic residues responsible for the formation of a small hydrophobic core which offers additional stabilization of the zinc finger domain. Panel (a) shows the alignment of the TFIIIA-like zinc finger domains from different organisms. The green color denotes residues that are responsible for the hydrophobic core formation in most CCHH zinc fingers (L17, F11, and L2). Yellow and blue indicate the coordinating Cys and His residues, respectively. Panel (b) shows the 3D NMR structure of 15-th ZF from zinc finger protein 478 [PDB: 2YRH] Figure \(17\) shows an interactive iCn3D model of C2H2-type zinc finger domain (699-729) from zinc finger protein 473 (2YRH). Overall, zinc finger motifs display considerable versatility in binding modes, even between members of the same class (e.g., some bind DNA, others protein), suggesting that they are stable scaffolds that have evolved specialized functions. For example, zinc finger-containing proteins function in gene transcription, translation, mRNA trafficking, cytoskeleton organization, epithelial development, cell adhesion, protein folding, chromatin remodeling, and zinc sensing, to name but a few. Zinc-binding motifs are stable structures, and they rarely undergo conformational changes upon binding their target. The last binding domain that we will consider in detail here is the helix-loop-helix domain found in Leucine zipper-containing proteins. Specifically, bZIPs (Basic-region leucine zippers) are a class of eukaryotic transcription factors. The bZIP domain is 60 to 80 amino acids in length with a highly conserved DNA binding basic region and a more diversified leucine zipper dimerization region. The two regions form α-helical structures that are connected via a looped region. This forms a core helix-loop-helix (HLH) structure within each monomer of the protein. Two monomers then join through the formation of a leucine zipper junction forming a heterodimeric protein structure. The resulting heterodimer can bind with DNA in a sequence-specific manner through the basic α-helices as shown in Figure \(18\). Specifically, basic residues, such as lysines and arginines, interact in the major groove of the DNA, forming sequence-specific interactions ). Most bZIP proteins show a high binding affinity for the ACGT motifs. The bZIP heterodimers exist in a variety of eukaryotes and are more common in organisms with higher evolution complexity. Figure \(19\) shows an interactive iCn3D model of the GCN4 basic region leucine zipper binds DNA as a dimer of uninterrupted alpha helices (1YSA). Epigenetics and Transgenerational Inheritance Even though all somatic cells of a multicellular organism have the same genome, different cell types have different transcriptomes (sets of all expressed RNA molecules), different proteomes (sets of all proteins), and, hence, different functions. Cell differentiation during embryonic development requires the activation and repression of specific sets of genes by the action of cell lineage-defining transcription factors. Within a cell lineage, gene activity states are often maintained over several rounds of cell divisions (a phenomenon called “cellular memory” or “cellular inheritance”). Since the rediscovery of epigenetics some 30 years ago (it was originally proposed by Conrad Hal Waddington in the early 1940s), cellular inheritance has been attributed to gene regulatory feedback loops, chromatin modifications (DNA methylation and histone modifications) as well as long-lived non-coding RNA molecules, which collectively are called the “epigenome”. Among the different chromatin modifications, DNA methylation and polycomb-mediated silencing are probably the most stable ones and endow genomes with the ability to impose silencing of transcription of specific sequences even in the presence of all of the factors required for their expression. Defining Transgenerational Epigenetic Inheritance The metastability of the epigenome explains why development is both plastic and canalized, as originally proposed by Waddington. Although epigenetics deal only with the cellular inheritance of chromatin and gene expression states, it has been proposed that epigenetic features could also be transmitted through the germline and persist in subsequent generations. The widespread interest in “transgenerational epigenetic inheritance” is nourished by the hope that epigenetic mechanisms might provide a basis for the inheritance of acquired traits. Yes, Lamarck has never been dead and every so often raises his head, this time with the help of epigenetics. Although acquired traits concerning body or brain functions can be written down in the epigenome of a cell, they cannot easily be transmitted from one generation to the next. For this to occur, these epigenetic changes would have to manifest in the germ cells as well, which in mammals are separated from somatic cells by the so-called Weismann barrier. Further, the chromatin is extensively reshaped during germ cell differentiation as well as during the development of totipotent cells after fertilization, even though some loci appear to escape epigenetic reprogramming in the germline. Long-lived RNA molecules appear to be less affected by these barriers and therefore more likely to carry epigenetic information across generations, although the mechanisms are largely unsolved. Evidence for Transgenerational Epigenetic Inheritance In the past 10 years, numerous reports on transgenerational responses to environmental or metabolic factors in mice and rats have been published. The factors include endocrine disruptors, high-fat diet, obesity, diabetes, undernourishment as well as trauma. These studies investigated DNA methylation, sperm RNA, or both. For example, when male mice are made prediabetic by treatment with streptozotocin it affects the DNA methylation patterns in their resulting sperm, as well as the pancreatic islets of F1 and F2 of the resulting offspring. Furthermore, studies have shown that traumatic stress in early life altered behavioral and metabolic processes in the progeny and that injection of sperm RNAs from traumatized males into fertilized wild-type oocytes reproduced the alterations in the resulting offspring. In humans, epidemiological studies have linked food supply in the grandparental generation to health outcomes in the grandchildren. An indirect study based on DNA methylation and polymorphism analyses has suggested that sporadic imprinting defects in Prader–Willi syndrome are due to the inheritance of a grandmaternal methylation imprint through the male germline. Because of the uniqueness of these human cohorts, these findings still await independent replication. Most cases of segregation of abnormal DNA methylation patterns in families with rare diseases, however, turned out to be caused by an underlying genetic variant. Thus, studies of this nature must rule out the effects of traditional genetic inheritance as being a factor of the observed phenotypes. Genetic inheritance alone cannot fully explain why we resemble our parents. In addition to genes, we inherited from our parents the environment and culture, which in parts have been constructed by the previous generations as shown in Figure \(20\). A specific form of the environment is our mother’s womb, to which we were exposed during the first 9 months of our life. The maternal environment can have long-lasting effects on our health. In the Dutch hunger winter, for example, severe undernourishment affected pregnant women, their unborn offspring, and the offspring’s fetal germ cells. The increased incidence of cardiovascular and metabolic disease observed in F1 adults is not due to the transmission of epigenetic information through the maternal germline, but a direct consequence of the exposure in utero, a phenomenon called “fetal programming” or—if fetal germ cells and F2 offspring are affected—“intergenerational inheritance”. Panel shows that offspring inherit from their parent's genes (black), the environment (green), and culture (blue). Genes and the environment affect the epigenome (magenta) and the phenotype. Culture also affects the phenotype, but at present, there is no evidence of a direct effect of culture on the epigenome (broken blue lines). It is a matter of debate, how much epigenetic information is inherited through the germline (broken magenta lines). G genetic variant, E epigenetic variant. Panel b shows that an epimutation (promoter methylation and silencing of gene B in this example) often results from aberrant read-through transcription from a mutant neighboring gene, either in sense orientation as shown here or in antisense orientation. The presence of such a secondary epimutation in several generations of a family mimics transgenerational epigenetic inheritance, although it represents genetic inheritance. Black arrow, transcription; black vertical bar, transcription termination signal; broken arrow, read-through transcription Roadmap to Proving Transgenerational Epigenetic Inheritance Here are some steps to show that inheritance is determined by epigenetics and not classical genetics. • Rule out genetic, ecological, and cultural inheritance. For studies in mice and rats, inbred strains and strictly controlled environments need to be used. When a pregnant female animal is exposed to a specific environmental stimulus, F3 offspring and subsequent generations must be studied to exclude a direct effect of the stimulus on the embryos’ somatic cells and germ cells. Even more desirable is the use of in vitro fertilization (IVF), embryo transfer, and foster mothers. When a male animal is exposed to an environmental stimulus, F2 offspring must be studied to exclude transient effects on germ cells. To ensure that any phenotype is exclusively transmitted via gametes, IVF must be used, controlling for possible artifacts relating to IVF. In contrast with laboratory animals, it is impossible to rule out ecological and cultural inheritance in humans, but genetic effects should and can be excluded. If an epimutation follows Mendelian inheritance patterns, be cautious: you are more likely looking at a secondary epimutation and genetic inheritance. Study the haplotype background of the epimutation: if in a given family it is always on the same haplotype, you are again most likely dealing with a secondary epimutation. Do whole genome sequencing to search for a genetic variant that might have caused the epimutation and be aware that this variant might be distantly located. Good spots to start looking at are the two neighboring genes, where a mutation might cause transcriptional read-through in sense or antisense orientation into the locus under investigation. Unfortunately, if you don’t find anything, you still cannot be 100% sure that a genetic variant does not exist. • Identify the responsible epigenetic factor in the germ cells. Admittedly, this is easier said than done, especially in female germ cells, which are scarce or unavailable. Be aware that germ cell preparations may be contaminated with somatic cells or somatic DNA. Use swim-up (sperm) or micromanipulation techniques to purify germ cells to the highest purity. Exclude the presence of somatic cells and somatic DNA by molecular testing, for example by methylation analysis of imprinted genes, which are fully methylated or fully unmethylated only in germ cells. • Demonstrate that the epigenetic factor in the germ cells is responsible for the phenotypic effect in the next generation. If possible, remove the factor from the affected germ cells and demonstrate that the effect is lost. Add the factor to control germ cells and demonstrate that the effect is gained. While RNA molecules can and have been extracted from the sperm of exposed animals and injected into control zygotes, DNA methylation, and histone modifications cannot easily be manipulated (although CRISPR/Cas9-based epigenome editors are being developed and used for this purpose), and all of these experiments can hardly be done in humans. In light of these problems, this might currently be too much to ask for to prove transgenerational epigenetic inheritance in humans, but should, nevertheless, be kept in mind and discussed.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/03%3A_Unit_III-_Information_Pathway/27%3A_Regulation_of_Gene_Expression/27.02%3A_Regulation_of_Gene_Expression_in_Eukaryotes.txt

These questions derive from the Research Literature Module - Carbon Capture Using Carbonic Anhydrase Question $1$ Using the equation below, at what ratio of CO2/[HCO3-] would the rate for the forward reaction (CO2 sequestration) be cut in half? v_0=\frac{V_M S}{K_M\left(1+\frac{I}{K is}\right)+S} If you need some help, hover over - Give me a hint! Answer \begin{gathered} v_{-I}=\frac{V_M S}{K_M+S}=\frac{(1)(1)}{(1+1)}=0.5 \ v_{+I}=\frac{V_M S}{K_M\left(1+\frac{[I]}{K_{i s}}\right)+S}=\frac{(1)(1)}{1\left(1+\frac{[I]}{2}\right)}=0.25=\frac{1}{1+\frac{[I]}{2}} \ 0.25\left(1+\frac{[I]}{2}\right)=1 \ 1+\frac{[I]}{2}=4 \ \frac{[I]}{2}=3 \ {[I]=6} \end{gathered} Hence it doesn't take much HCO3- buildup to inhibit the "capture" of CO2! Carbonic Anhydrase - Mechanism These questions derive from the Research Literature Module - Carbon Capture Using Carbonic Anhydrase Carbonic Anhydrase - Structure and Mechanism 1 An active site Zn2+ appears to bind a water molecule and reduce its pKa such that the bound form is OH-. This is illustrated in the left panel of Figure $1$ below, which depicts the local environment of the bound Zn2+ (coordinated by histidine side chains and an OH-) in the absence (left) and presence (right) of CO2. Note that the back histidine is difficult to barely visible (but still evident) in both structures. To assist in viewing the structure, the right panel shows an interactive iCn3D model of Zn- human carbonic anhydrase II at pH 7.8 and 0 atm CO (6LUW). Figure $1$: Left panel: Coordination of OH- to Zn2+ in carbonic anhydrase in the absence (left) and presence (right) of substrate CO2. Right panel: Zn- human carbonic anhydrase II at pH 7.8 and 0 atm CO (6LUW) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...43DYyZFvHJpZn9 Question $\PageIndex1}$ What is the coordination geometry of the Zn ion? Answer tetrahedral Question $2$ Draw a simplified reaction mechanism showing bicarbonate formation from the two reactants, CO2 and OH-. Answer The enzyme is reversible and in humans is important in CO2 transport in respiration and maintaining intracellular pH, which is also its key role in most organisms. CO2, like the other atmospheric gases O2 and N2, are nonpolar and have limited solubility in water. The solubility of these gases in water at 20oC and 1 atm pressure, in g/L and mM, are shown in Table $1$ below. Gas (ref) solubility (aq) (g/L) solubility (mM) CO2 1.7 38 O2 0.044 1.3 N2 0.019 0.68 Question $3$ Offer reasons that explain the significantly higher (but still low) solubility of CO2 in water compared to O2 and N2. Answer CO2 is considered a nonpolar molecule since it has no net molecular dipole. However, it does have 2 bond dipoles (pointing in opposite directions), so the carbon atom is δ+ while the Os are δ-. This probably contributes to its greater solubility than N2 and O2 which don't have bond dipoles. CO2 would not orient itself in a dipole electric field, but it would to some extent in a quadrupole (4 poles) electric field where the positive potentials are oriented north and south and the negative potentials at east and west. CO2 has a quadrupole moment. In addition, the continued reaction of CO2 and water to form the weak acid carbonic acid would contribute to its higher apparent solubility. It is not clear to the authors if these contributions to solubility are accounted for in the experimental values of solubility. A quadruple and its associated magnetic field with oriented CO2. https://commons.wikimedia.org/wiki/F...quadrupole.svg Question $4$ How does the enzyme facilitate the transport of CO2 in blood? How does it maintain intracellular pH? Answer It converts the poorly soluble carbon unit in CO2 to the strongly soluble bicarbonate anion. HCO3-. A simple explanation for maintaining intracellular pH comes from the chemical equation below. CO2(aq) + H2O (l) ↔ H2CO3 (aq) + H2O(l) H3O+(aq) + HCO3-(aq). HCO3- is the conjugate base of the weak acid, H2CO3 so the system is a classic buffer. For a complete explanation of why the system can act as a buffer at neutral pH even though the pKa of the weak acid is 3.6, see Chapter 2.3 for review. Let's explore the structure of two different CAs, human carbonic anhydrase II and the carbonic anhydrase from Neisseria gonorrhea. Human carbonic anhydrase II The structure of native human carbonic anhydrase II and its catalytic mechanism is shown in Figure $5$ below. Figure $5$: Structure of native human carbonic anhydrase II (Zn-CA II) and its catalytic mechanism. Kim, J.K., Lee, C., Lim, S.W. et al.Nat Commun 11, 4557 (2020). https://doi.org/10.1038/s41467-020-18425-5. Creative Commons Attribution 4.0 International License, http://creativecommons.org/licenses/by/4.0/. Panel a shows the active site consists of the zinc binding site, hydrophobic/hydrophilic regions, and the entrance conduit (EC). Panel b shows the water networks in the active site that are responsible for the proton transfer (red) and substrate/product/water exchange (blue) during enzyme catalysis. Panel c shows the forward reaction mechanism of Zn-CA II. The active site itself lies at the bottom of a deep cavity (15 Å deep) in the protein, which is readily accessible to solvent An interactive iCn3D model of human carbonic anhydrase II with bound bicarbonate and CO2 (2VVB) is shown in Figure $6$ below. Figure $6$: Human carbonic anhydrase II with bound bicarbonate and CO2 (2VVB) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...ibgmgsm3UWhjX6 The active site residues shown in Figure 6 are labeled and shown as sticks. Bound CO2 and HCO3- are also shown as sticks. Question $5$ This question addresses the Biomolecular Visualization Framework theme(s) Molecular Interactions (MI), Atomic Geometry (AG) Using iCn3D to show the noncovalent interactions between bicarbonate, Zn, and the protein by using iCn3D. Measure the distance between the hydrogen-bonded atoms in HCO3- and Thr 199. iCn3D instructions Trackpad and Mouse Controls rotate: click and drag (mouse: left click and drag) zoom: pinch and spread (mouse: rotate the scroll wheel) translate: two-finger click and drag (mouse: right click and drag) Re-center: left click View from the top menu bar, then select “Center Selection” •Note: ctrl-click on a PC = command-click on Mac; alt-click on PC = option click on Mac 1. Open the external link: https://structure.ncbi.nlm.nih.gov/i...ibgmgsm3UWhjX6 2. From the top menu bar, choose Analysis, Interactions 3. In the new popup window select the following prompts, then click 4. 3D Display interactions 4. Close all but the main modeling window. 5. Zoom into the bicarbonate binding site. From the top menu bar, choose Analysis, Distance, distance between 2 atoms, and pick the two atoms (by holding down the Alt key or Option on a Mac) involved in the hydrogen bond between the bicarbonate and the amide 6. Rescale the label size by choosing Analysis, Label Scale, 0.4. 7. From the top menu bar choose Select, Toggle highlights to remove yellow and box highligting. 8. Save a PNG file by choosing 9. You can reload the PNG file directly into iCn3D by choosing File, Open File, iCn3D PNG image Answer The blue dotted lines are ion-ion interactions. Note the green hydrogen bond between T199 and the bound bicarbonate. Remember that hydrogen atoms are not shown in PDB files from x-ray structures. The H-bond distance is 3.1 angstroms. For some reason, the H-bond does not show unless the initial constraint distances are moved to 4.2 Angstroms. Question $6$ What is a likely function of Val, Leu and Trp cluster in CAII (shown in Figures 5 and 6)? Answer These side chains are all hydrophobic (as illustrated in gold in Figure 5) and provide a weak binding environment for nonpolar CO2. Question $7$ What ligand would likely replace OH- at low pH values? What would happen to the activity of the enzyme at lower pHs? Answer At low pH, ie. at pH values lower than the pKa of the Zn-bound water (which deprotonates to form the OH- ligand and nucleophile), the ligand and nucleophile would be H2O. The enzyme would display a lower activity given the weaker nucleophile. Question $8$ From the mechanism shown in Figure 5, does bicarbonate coordinate the Zn2+ ion in a monodentate or bidentate manner? Answer Monodentate as only 1 coordinate covalent bond forms on an electron pair donation from the bicarbonate to the Zn2+. Question $9$ Write a verbal description of the mechanism of CAII based on Figure 5 Answer CO2 binds to the active site through loose association with the cluster of hydrophobic side chains. The Zn2+ bound OH- acting as a nucleophile attacks the central carbon of the CO2 forming HCO3. The carbonate forms a monodentate interaction with Zn and also a hydrogen bond to Thr 199. The HCO3- is then displaced by an incoming water molecule. The other product of the reaction, H+, moves through a hydrogen bond network of water molecules (W1 and W2) to His 64 and eventually to bulk water. The interactions with substrate and products are weak allowing fast exchange. Question $10$ Why is a proton transfer path needed? Answer H+ is a product of the reaction: CO2 (g) + H2O ↔ H2CO3 (aq) ↔ HCO3- (aq) + H+ (aq). It must depart to prevent charge build-up, maintain charge balance, and keep the correct electrostatic environment of the active site. Question $11$ Thr 199 plays a key role in the mechanism. State a reason for its importance in the reversible reaction. Answer Thr 199 supplies two hydrogen bonds to the bicarbonate. It actually destabilizes bicarbonate bonding with respect to a T199A mutant. In the mutant, carbonate might bind Zn2+ in a bidentate fashion, leading to tighter binding and a slower dissociation rate of the product, HCO3-. Question $12$ The dissociation constant KD (or Kis) for bicarbonate binding to HCAII is about 77 mM. What kind of inhibitor might it be for the forward reaction? Answer Given that it binds in the active site and would prevent binding of the substrate for the forward reaction, CO2, it is a competitive inhibitor. Question $13$ Using the equation below, at what ratio of CO2/[HCO3-] would the rate for the forward reaction (CO2 sequestration) be cut in half? Assume the VM=1, KM forward reaction is 1, [S] = 1 and Kis is 2 (a wide range of values are reported in the Brenda Database. v_0=\frac{V_M S}{K_M\left(1+\frac{I}{K is}\right)+S} If you need some help, hover over - Give me a hint! Answer \begin{gathered} v_{-I}=\frac{V_M S}{K_M+S}=\frac{(1)(1)}{(1+1)}=0.5 \ v_{+I}=\frac{V_M S}{K_M\left(1+\frac{[I]}{K_{i s}}\right)+S}=\frac{(1)(1)}{1\left(1+\frac{[I]}{2}\right)}=0.25=\frac{1}{1+\frac{[I]}{2}} \ 0.25\left(1+\frac{[I]}{2}\right)=1 \ 1+\frac{[I]}{2}=4 \ \frac{[I]}{2}=3 \ {[I]=6} \end{gathered} Hence it doesn't take much HCO3- buildup to inhibit the "capture" of CO2! Question $14$ The rate-limiting step for human CA II is the dissociation of a proton from Zn2+-bound water and not the removal of the resulting proton from the enzyme. What does that imply about the rate of removal of the proton from the enzyme? Answer It must be very fast, that is at diffusion-controlled limits through the H-bond channel. Synthetic mimetics of the active site of CA have been made. These are heteromacrocycles (similar to the heme of hemoglobin) as shown in Figure $9$ below. Figure $9$: The macrocycle mimetic has three imidazole groups coordinating zinc. Question $15$ The macrocycle mimetic has three imidazole groups coordinating zinc. Is the bicarbonate coordinated to the Zn2+ in a monodentate or bidentate fashion? From the "denticities" of the interactions of bicarbonate and Zn2+ for human CAII and the mimetic, which catalyst, CAII or the macrocycle would you expect to have a lower KD for bicarbonate? How might this affect the rate-limiting step for the mimetic? Answer The mimetic is bidentate, is it should bind more tightly to bicarbonate, hindering its dissociation, and hence making "product" inhibition more likely. Question $16$ What is the utility of having both CO2 and HCO3- bind weakly to the enzyme Answer Weak binding "permits their rapid exchange. The hydrogen-bonding arrangement in the active site is such that the water or hydroxide ion donates a hydrogen bond to a proximal threonine (Thr199 in hCA II) because the hydroxyl group of this residue is forced to donate its hydrogen in a hydrogen bond to a negatively charged glutamate side chain (Glu106 in hCA II). Site-directed mutations have confirmed this model of the catalytic mechanism. The substrates/products carbon dioxide/bicarbonate are fairly weakly bound against a hydrophobic wall in the active site, which permits their rapid exchange."

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/Enzyme_Kinetics_Problems/Carbonic_Anydrase_Inhibition.txt

Introduction Instead of presenting a litany of end-of-chapter or end-of-book questions that are not linked in content or concepts, we will present a number of problem-solving assessments linked to research literature that deal with key challenges that face the world today. We will call these research literature modules (RLM). Each will focus on a particular biological system (enzyme, pathways, etc) and contain a series of sequential and linked questions on a particular protein, for example, and its function. The problems are summative and hence require an understanding of structure, noncovalent interactions, binding, kinetics, and reaction mechanisms. The modules are guided and have elements of problem-solving and POGIL questions. Each Module will ultimately focus on the structure and properties of key biomolecules. The modules will: 1. Address ASBMB Core concepts and Learning Objectives (generalized below) through analysis and interpretation of research findings in one or more research publications. • Energy is required by and transformed in biological systems. • Macromolecular structure determines function and regulation. • Information storage and flow are dynamic and interactive. • Biochemical systems maintain a state of homeostasis, a steady stable state while continually adjusting to conditions, which requires energy input, organization, and control mechanisms. • Evolution plays a pervasive role in shaping the form and function of all biological molecules and organisms. 2. Link to critical problems facing the world (see the Table) which have clearly identified biochemical components. These critical problems include health care disparities, climate change, pandemics, addiction, childhood trauma, food insecurity, biodiversity loss, ecosystem (ocean, soil, forest) health, and misinformation/disinformation. These problems are often linked and not mutually exclusive. Most biochemistry textbooks focus on problems using biomedical examples. Expanding to study key world problems that are not directly biomedical and are underrepresented in textbooks, allows students to apply their acquired knowledge and understanding into different areas. 3. Follow general features found in problem-based learning and in case studies, which provide contextual applications for the detailed learning opportunities found in biochemistry books and courses. • A broad introduction (text, videos, personal narratives) describing the critical world problem and the relevancy of the selected biochemical system to the problem • A more detailed description of the selected biochemical system, including links to specific locations in Fundamentals of Biochemistry as well as external resources • Research literature results (graphs, tables, models, etc), taken from journals that allow derivatives and reuse by appropriate Creative Commons licensing (for example, CC BY 4.0), for interpretation 4. Focus on representative biomacromolecules (protein, nucleic acid, glycan, lipid and combination of them) relevant to the broader problem for which detailed structure/function questions can be explored 5. Explicitly address and link to appropriate BioMolViz framework themes, goals and objectives to the biomolecules key to the RLM. Relationship of RLMs to BioMolViz and Molecular CaseNet The completed RLMs will consist of a broad introduction and relevant biochemical research findings woven into a narrative that will include nested questions based on the literature with an ultimate focus on a key biomacromolecule. It will not take the form or detail of a full case study as found in Molecular CaseNet (headed by Shuchi Dutta and its Steering Committee, which includes Henry Jakubowski, who is also on the Steering Committee of BioMolViz ). As the RLMs in Fundamentals of Biochemistry and indeed the whole text, as well as the Molecular CaseNet are free online educational resources (OERs), both communities can freely share resources. Since the RLMs have some attributes of case studies, we hope that contributors to Molecular CaseNet will freely use the RLMs and convert them to more expansive case studies, housed within Molecular CaseNet. Likewise, the research literature-based questions in the RLMs that focus on biomacromolecule structures will be explicitly linked to the themes, goals and objectives of the BioMolViz literacy framework. However, the specific questions will not be included in the web repository created by BioMolViz. The repository questions have gone through many iterative cycles of construction, revision, external review by expert panels, and validation by actual classroom use. Instead, the questions in Fundamentals of Biochemistry RLMs that target specific biomolecular visualization framework objectives will help to expand knowledge and understanding of biomolecule visual literacy and BioMolViz objectives, which ultimately is the goal of BioMolViz. A full semester of biochemistry would be necessary to complete a full RLM, as the questions extend from structure, binding, kinetics, mechanism, metabolism, and signal transduction. Yet parts of a complete RLM could be completed after students complete the corresponding chapter in the book. Hence parts of a given RLM will be listed in Volume 5 under the corresponding topic (carbohydrate structure, for example). A link will be provided back to the home RLM from which the questions were derived World Challenges as the Bases for the Research Literature Modules Here are the world challenges we have selected that we serve as the bases for the RLMs. World Problems Research Literature Modules Health Disparities Type II Diabetes Orphan receptors Poverty and stress response: Poverty and epigenetics PM2.5s Pb pollution Pollution (air/water) Climate Change Thermal tolerance plants Carbon Capture Photosynthesis, CO2 sequestration Heat Stroke Biofuels Modeling climate change Pandemics Vaccine Development Ebola Malaria Emerging Diseases Evolution Addiction Natural/Synthetic opiates Alcohol abuse Trauma PTSD Food Insecurity photosynthesis fertilizers Loss of Biodiversity extinction Ecosystem Health Soil Oceans Forest Mis- Disinformation Western blots and image modifications

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/Global_Challenges_-_Literature-Based_Guided_Assessments_(LGAs)/1.__Global_Challenges%3A_Literature-based_Guided.txt

Research Literature Module - Carbon Capture Using Carbonic Anhydrase Critical World Challenges Climate Change Key Words, Concepts: protein structure, structure/function relationships, enzyme kinetics, enzyme mechanisms, reaction mechanisms, Western blot analysis, site-directed mutagenesis, biomolecular visualization, computational modeling, graphic analysis The Problem Our climate is changing as planetary temperatures rise from increasing amounts of the greenhouse gas carbon dioxide released into the atmosphere (detailed in Chapter 31) on the burning of fossil fuels. The rate of release is unparalleled in geological history. Present levels (415 ppm) have not been seen for at least 3 million years. Human societies and cultures have had the opportunity to develop in relatively stable climatic conditions. Figure $1$ below shows the rise in atmospheric CO2 over the last 1000 years. Figure $1$: CO2 levels in the atmosphere over the last 1000 years. Our world in data. https://ourworldindata.org/ The steep rise around 1790 coincides with the start of the industrial revolution. The rise in atmospheric CO2 has led to a corresponding rise in the average global temperatures, as illustrated in Figure $2$ below. Figure $2$: Average global temperature changes over the last 1000 years. Our world in data. https://ourworldindata.org/ The per capita emissions of CO2 across the world derive from the use of coal, oil, and gas, as illustrated in Figure $3$ below. Figure $3$: Per capita emission of CO2 from fossil fuel type If we want to decrease emissions, we also need to know in which economic sectors fossil fuels are used. The main sources of global energy-related CO2 emissions by sector are shown in Figure $4$: Figure $4$: Global energy-related CO2 emissions by sector. Updated on 10/26/22. https://www.iea.org/data-and-statist...ions-by-sector. IEA. License: CC BY 4.0 Note the use of coal, gas, and oil for energy production (electricity) accounts for 40% of global CO2 emissions, with transportation (mostly through the use of gasoline and diesel fuel) and industry accounting for about 25% each. Simple chemistry tells us that there are two ways to decrease the amount of product (in this case CO2) in a chemical reaction: • decrease the concentration of reactants (i.e. reduce fossil fuel use) • remove the product, in this case, CO2 from the air. The latter process is called carbon capture or sequestration. It is a daunting process that nature has mastered (through photosynthesis), but it clearly can't keep up with the huge injection of CO2 in the atmosphere caused by burning fossil fuel. We simply can't stop using fossil fuels, which would result in huge economic and social unrest. Alternative green fuels (solar, wind, for example) are being rapidly expanded but can't replace fossil fuels for many years. One of the reasons is that fossil fuels are very energy-dense (MJ/kg) compared to other sources of energy. It's also fascinating to look at the energy transitions humans have made over time. Figure $5$ below shows the energy transition over a log-time scale (for presentation purposes) as well as the energy densities of individual sources. Figure $5$: Human-created energy transitions (log time scale) and energy densities of individual sources New technologies are needed to capture CO2. We have to move much faster in a new clean energy transition than we have in our entire history. A potentially ideal solution would be to capture CO2 from power plants before they reach the atmosphere. We will now look at research into an old enzyme, carbonic anhydrase, that is being repurposed for industrial-level carbon capture, carbonic anhydrase. Carbonic Anhydrase (CA) We have already encountered this enzyme before (Chapter 6.1). It catalyzes the hydration of CO2 (g) as shown below. CO2 (g) + H2O ↔ H2CO3 (aq) ↔ HCO3- (aq) + H+ (aq) It is among the fastest of all enzymes, with a kcat of 106 s-1 and a kcat/Km of 8.3 x 107 M-1s-1 (reference). It is diffusion controlled in that the rate of diffusion of reactants and products, not the chemical steps, determine the reaction rate. It can convert 106 molecules of CO2(g) to HCO3- each second. No wonder scientists and engineers are studying it to capture CO2. It's a big challenge though to capture CO2 released on combustion of coal or natural gas in a power plant. Here are two problems that must be overcome: • The enzyme must be thermostable at elevated temperatures to capture the CO2 found in high-temperature power plant emissions • The enzyme is reversible so it will be inhibited by the product HCO3- • The enzyme must be stable to somewhat alkaline conditions (pH of 0.1M NaHCO3 = 8.3) For carbon capture from fossil fuel emissions, CA is immobilized by surface adsorption, covalent attachment, encapsulation, and entanglement. Immobilized enzymes are typically more thermostable and can be used in flow-through as opposed to solution phase capture. The immobilized enzyme matrix must withstand high temperatures (up to 100°C, and alkaline solvents used to strip the matrix for reuse. The enzyme is found throughout life and typically has an active site Zn2+. There are 8 families, α, β, γ, δ, ζ η, θ, and ι, with the α family being the most abundant. The α forms are generally active as dimers, but can act as monomers and tetramers.. There are 15 isoforms of the α form in humans and have a prime role in pH regulation. They are found in bacteria, fungi, plants, and algae. β-CAs are found in some types of bacteria, archaea, fungi, some higher plants, and invertebrates. CA in chloroplasts (and mitochondria (algae) are involved in carbon fixation. We will focus our attention on engineering carbonic anhydrase to make them more thermostable, alkali insensitive, and less susceptible to product inhibition by bicarbonate. Natural enzymes can be isolated and selected for thermal and alkali stability. In addition, new versions selected for these properties can be engineered using directed evolution or site-directed mutagenesis. You wish to increase the thermal stability of a protein using mutagenesis. Essentially you wish to perturb the equilibrium between the folded (native) protein and the unfolded (denatured) protein so as to preferentially stabilized the native state. Question $1$ Using mutagenesis, what residues might you change in a native protein to make it more stable at higher temperatures? Answer A characteristic of the native state of the protein is its conformational stability compared to the conformational flexibility of the many possible denatured states. In addition, the protein must undergo conformational changes as it unfolds. Hence anything that restricts conformational flexibility might preferentially stabilize the native state. These would include changing single or pairs of side chains to allow the formation of more salt bridges and intrachain disulfide bonds, as well as hydrogen bonds. Loops with greater flexibility, as determined by B-factors in the crystal structure files, or by molecular dynamic simulations, could be changed to contain a disulfide, which would clearly stabilize a flexible loop. Question $2$ What measurements would you make to quantitate the change in thermal stability? Answer Measures a signal that changes with increasing temperature. The signal can be enzyme activity, or more easily a spectroscopic signal such as absorbance at 280 nm or fluorescence as a function of temperature. Alternatively, the stability at room temperature could be measured using urea as a perturbant. These are discussed in Chapter 4.12. The actual amino acid composition and more strangely specific dipeptide sequences within a sequence are associated with thermal stability of hyperthermophilic proteins. For example, proteins from two different types of archaea with different optimal growth temperatures show that the one with the higher growth temperature have significantly higher levels of VK, KI, YK, IK, KV, KY, and EV and decreased levels of DA, AD, TD, DD, DT, HD, DH, DR, and DG. Similar experiments have been done in bacterial cells. Using machine learning, the dipeptide sequences KH, KR, TF, PM, F∗∗N, V∗∗Y, MW, and WQ were important in themostability where the * denotes a gap in the residues. Structure and Mechanism An active site Zn2+ appears to bind a water molecule and reduce its pKa such that the bound form is OH-. This is illustrated in the left panel of Figure $6$ below, which depicts the local environment of the bound Zn2+ (coordinated byhistidine side chains and an OH-) in the absence (left) and presence (right) of CO2. Note that the back histidine is difficult to barely visible (but still evident) in both structures. To assist in viewing the structure, the right panel shows an interactive iCn3D model of Zn- human carbonic anhydrase II at pH 7.8 and 0 atm CO (6LUW). Figure $6$: Left panel: Coordination of OH- to Zn2+ in carbonic anhydrase in the absence (left) and presence (right) of substrate CO2. Right panel: Zn- human carbonic anhydrase II at pH 7.8 and 0 atm CO (6LUW) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...43DYyZFvHJpZn9 Question $3$ What is the coordination geometry of the Zn ion? Answer tetrahedal Question $4$ Draw a simplified reaction mechanism showing bicarbonate formation from the two reactants, CO2 and OH-. Answer The enzyme is reversible and in humans is important in CO2 transport in respiration and maintaining intracellular pH, which is also its key role in most organisms. CO2, like the other atmospheric gases O2 and N2, are nonpolar and have limited solubility in water. The solubility of these gases in water at 20oC and 1 atm pressure, in g/L and mM, are shown in Table $1$ below. Gas (ref) solubility (aq) (g/L) solubility (mM) CO2 1.7 38 O2 0.044 1.3 N2 0.019 0.68 Question $5$ Offer reasons that explain the significantly higher (but still low) solubility of CO2 in water compared to O2 and N2. Answer CO2 is considered a nonpolar molecule since it has no net molecular dipole. However, it does have 2 bond dipoles (pointing in opposite directions), so the carbon atom is δ+ while the Os are δ-. This probably contributes to its greater solubility than N2 and O2 which don't have bond dipoles. CO2 would not orient itself in a dipole electric field, but it would to some extent in a quadrupole (4 poles) electric field where the positive potentials are oriented north and south and the negative potentials at east and west. CO2 has a quadrupole moment. In addition, the continued reaction of CO2 and water to form the weak acid carbonic acid would contribute to its higher apparent solubility. It is not clear to the authors if these contributions to solubility are accounted for in the experimental values of solubility. A quadruple and its associated magnetic field with oriented CO2. https://commons.wikimedia.org/wiki/F...quadrupole.svg Question $6$ How does the enzyme facilitate the transport of CO2 in blood? How does it maintain intracellular pH? Answer It converts the poorly soluble carbon unit in CO2 to the strongly soluble bicarbonate anion. HCO3-. A simple explanation for maintaining intracellular pH comes from the chemical equation below. CO2(aq) + H2O (l) ↔ H2CO3 (aq) + H2O(l) H3O+(aq) + HCO3-(aq). HCO3- is the conjugate base of the weak acid, H2CO3 so the system is a classic buffer. For a complete explanation of why the system can act as a buffer at neutral pH even though the pKa of the weak acid is 3.6, see Chapter 2.3 for review. We'll explore the structure of two different CAs, human carbonic anhydrase II and the carbonic anhydrase from Neisseria gonorrhea in this guided problem-solving module. Human carbonic anhydrase II The structure of native human carbonic anhydrase II and its catalytic mechanism is shown in Figure $7$ below. Figure $7$: Structure of native human carbonic anhydrase II (Zn-CA II) and its catalytic mechanism. Kim, J.K., Lee, C., Lim, S.W. et al.Nat Commun 11, 4557 (2020). https://doi.org/10.1038/s41467-020-18425-5. Creative Commons Attribution 4.0 International License, http://creativecommons.org/licenses/by/4.0/. Panel a shows the active site consists of the zincbinding site, hydrophobic/hydrophilic regions, and the entrance conduit (EC). Panel b shows the water networks in the active site that are responsible for the proton transfer (red) and substrate/product/water exchange (blue) during enzyme catalysis. Panel c shows the forward reaction mechanism of Zn-CA II. The active site itself lies at the bottom of a deep cavity (15 Å deep) in the protein, which is readily accessible to solvent An interactive iCn3D model of human carbonic anhydrase II with bound bicarbonate and CO2 (2VVB) is shown in Figure $8$ below. Figure $8$: Human carbonic anhydrase II with bound bicarbonate and CO2 (2VVB) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...ibgmgsm3UWhjX6 The active site residues shown in Figure 8 are labeled and shown as sticks. Bound CO2 and HCO3- are also shown as sticks. Question $7$ This question addresses the Biomolecular Visualization Framework theme(s) Molecular Interactions (MI), Atomic Geometry (AG) Using iCn3D to show the noncovalent interactions between bicarbonate, Zn, and the protein by using iCn3D. Measure the distance between the hydrogen-bonded atoms in HCO3- and Thr 199. iCn3D instructions Trackpad and Mouse Controls rotate: click and drag (mouse: left click and drag) zoom: pinch and spread (mouse: rotate the scroll wheel) translate: two-finger click and drag (mouse: right click and drag) Re-center: left click View from the top menu bar, then select “Center Selection” •Note: ctrl-click on a PC = command-click on Mac; alt-click on PC = option click on Mac 1. Open the external link: https://structure.ncbi.nlm.nih.gov/i...ibgmgsm3UWhjX6 2. From the top menu bar, choose Analysis, Interactions 3. In the new popup window select the following prompts, then click 4. 3D Display interactions 4. Close all but the main modeling window. 5. Zoom into the bicarbonate binding site. From the top menu bar, choose Analysis, Distance, distance between 2 atoms, and pick the two atoms (by holding down the Alt key or Option on a Mac) involved in the hydrogen bond between the bicarbonate and the amide 6. Rescale the label size by choosing Analysis, Label Scale, 0.4. 7. From the top menu bar choose Select, Toggle highlights to remove yellow and box highligting. 8. Save a PNG file by choosing 9. You can reload the PNG file directly into iCn3D by choosing File, Open File, iCn3D PNG image Answer The blue dotted lines are ion-ion interactions. Note the green hydrogen bond between T199 and the bound bicarbonate. Remember that hydrogen atoms are not shown in PDB files from x-ray structures. The H-bond distance is 3.1 angstroms. For some reason, the H-bond does not show unless the initial constraint distances are moved to 4.2 Angstroms. Question $8$ What is a likely function of Val, Leu and Trp cluster in CAII (shown in Figures 7 and 8)? Answer These side chains are all hydrophobic (as illustrated in gold in Figure 7) and provide a weak binding environment for nonpolar CO2. Question $\PageIndex{x}$ What ligand would likely replace OH- at low pH values? What would happen to the activity of the enzyme at lower pHs? Answer At low pH, ie. at pH values lower than the pKa of the Zn-bound water (which deprotonates to form the OH- ligand and nucleophile), the ligand and nucleophile would be H2O. The enzyme would display a lower activity given the weaker nucleophile. Question $9$ From the mechanism shown in Figure 7, does bicarbonate coordinate the Zn2+ ion in a monodentate or bidentate manner? Answer Monodentate as only 1 coordinate covalent bond forms on an electron pair donation from the bicarbonate to the Zn2+. Question $10$ Write a verbal description of the mechanism of CAII based on Figure 7 Answer CO2 binds to the active site through loose association with the cluster of hydrophobic side chains. The Zn2+ bound OH- acting as a nucleophile attacks the central carbon of the CO2 forming HCO3. The carbonate forms a monodentate interaction with Zn and also a hydrogen bond to Thr 199. The HCO3- is then displaced by an incoming water molecule. The other product of the reaction, H+, moves through a hydrogen bond network of water molecules (W1 and W2) to His 64 and eventually to bulk water. The interactions with substrate and products are weak allowing fast exchange. Question $11$ Why is a proton transfer path needed? Answer H+ is a product of the reaction: CO2 (g) + H2O ↔ H2CO3 (aq) ↔ HCO3- (aq) + H+ (aq). It must depart to prevent charge build-up, maintain charge balance, and keep the correct electrostatic environment of the active site. Question $12$ Thr 199 plays a key role in the mechanism. State a reason for its importance in the reversible reaction. Answer Thr 199 supplies two hydrogen bonds to the bicarbonate. It actually destabilizes bicarbonate bonding with respect to a T199A mutant. In the mutant, carbonate might bind Zn2+ in a bidentate fashion, leading to tighter binding and a slower dissociation rate of the product, HCO3-. Question $13$ The dissociation constant KD (or Kis) for bicarbonate binding to HCAII is about 77 mM. What kind of inhibitor might it be for the forward reaction? Answer Given that it binds in the active site and would prevent binding of the substrate for the forward reaction, CO2, it is a competitive inhibitor. Question $14$ Using the equation below, at what ratio of CO2/[HCO3-] would the rate for the forward reaction (CO2 sequestration) be cut in half? Assume the VM=1, KM forward reaction is 1, [S] = 1 and Kis is 2 (a wide range of values are reported in the Brenda Database. v_0=\frac{V_M S}{K_M\left(1+\frac{I}{K is}\right)+S} If you need some help, hover over - Give me a hint! Answer \begin{gathered} v_{-I}=\frac{V_M S}{K_M+S}=\frac{(1)(1)}{(1+1)}=0.5 \ v_{+I}=\frac{V_M S}{K_M\left(1+\frac{[I]}{K_{i s}}\right)+S}=\frac{(1)(1)}{1\left(1+\frac{[I]}{2}\right)}=0.25=\frac{1}{1+\frac{[I]}{2}} \ 0.25\left(1+\frac{[I]}{2}\right)=1 \ 1+\frac{[I]}{2}=4 \ \frac{[I]}{2}=3 \ {[I]=6} \end{gathered} Hence it doesn't take much HCO3- buildup to inhibit the "capture" of CO2! Question $15$ The rate-limiting step for human CA II is the dissociation of a proton from Zn2+-bound water and not the removal of the resulting proton from the enzyme. What does that imply about the rate of removal of the proton from the enzyme? Answer It must be very fast, that is at diffusion-controlled limits through the H-bond channel. Synthetic mimetics of the active site of CA have been made. These are heteromacrocycles (similar to the heme of hemoglobin) as shown in Figure $9$ below. Figure $9$: The macrocycle mimetic has three imidazole groups coordinating zinc. Question $16$ The macrocycle mimetic has three imidazole groups coordinating zinc. Is the bicarbonate coordinated to the Zn2+ in a monodentate or bidentate fashion? From the "denticities" of the interactions of bicarbonate and Zn2+ for human CAII and the mimetic, which catalyst, CAII or the macrocycle would you expect to have a lower KD for bicarbonate? How might this affect the rate-limiting step for the mimetic? Answer The mimetic is bidentate, is it should bind more tightly to bicarbonate, hindering its dissociation, and hence making "product" inhibition more likely. Question $17$ What is the utility of having both CO2 and HCO3- bind weakly to the enzyme Answer "permits their rapid exchange. The hydrogen-bonding arrangement in the active site is such that the water or hydroxide ion donates a hydrogen bond to a proximal threonine (Thr199 in hCA II) because the hydroxyl group of this residue is forced to donate its hydrogen in a hydrogen bond to a negatively charged glutamate side chain (Glu106 in hCA II). Site-directed mutations have confirmed this model of the catalytic mechanism. The substrates/products carbon dioxide/bicarbonate are fairly weakly bound against a hydrophobic wall in the active site, which permits their rapid exchange." Carbonic anhydrase from Neisseria gonorrhea (ngCA) Data from: Jo, B., Park, T., Park, H. et al. Engineering de novo disulfide bond in bacterial α-type carbonic anhydrase for thermostable carbon sequestration. Sci Rep 6, 29322 (2016). https://doi.org/10.1038/srep29322. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ Now that we understand the general chemistry, structure, and reaction mechanism of carbon anhydrase (at least the alpha human CAII form), let's explore efforts to engineer more thermostable variants. One example is the carbonic anhydrase from N. gonorrheas. This CA has been used as a target for mutagenesis to increase thermal stability of the enzyme, through the introduction of new disulfide bonds. Even though only about 35% of the amino acids are identical, the overall structures are similar. This is illustrated in Figure $10$ below. Figure $10$: Alignment of the carbonic anhydrase from Neisseria gonorrhea (NG-CA) magenta,1KOQ) and human CA II (cyan, 2VVB) The active site is mostly conserved compared to human CA II. The Zn2+ bound water has a pKa of around 6.5, compared to the value of 7.0 in human CA II. The hydrophobic patch (pocket) is similar, with Phe 93, Leu 153 and Tyr 72 in the NG-CA replacing Phe 95, Phe 176, Phe 70 in human CA II, respectively. The histidine ligands to Zn2+ are His92 (94), His94 (96), and His111 (119), where the numbers in parentheses represent Hu CA II. The proton removed from Zn2+ bound water is transferred to His 66 (64 in human CA II) and then to His 64. The single disulfide bond between 181 and C28 is shown in Figure $1$ below Figure $11$: Single disulfide bond between 181 and C28 in wild type Carbonic anhydrase from Neisseria gonorrhea Question $18$ This question addresses the Biomolecular Visualization Framework theme(s) Atomic Geometry (AG) Identify the correct torsion angles in Figure $\PageIndex{x}$ above. Verbal definitions of torsional angles in a peptide chain are listed below. The successive atoms after the Cα leading away from the backbone atoms are Xβ-Xγ-Xδ-Xε (in that order). C is the backbone carbonyl C and N is the backbone nitrogen atom. • phi (φ) is the angle of right-handed rotation around N-Cα bond. φ = 0 if the Cα-C bond is cis (eclipsed) to the C-N bond. Values range from -180 to 180 degrees. • psi (ψ) is the angle of right-handed rotation around Cα -C bond. ψ = 0 if the C-N bond is cis (eclipsed) to the N-Cα bond. Values range from -180 to 180 degrees. • chi11) is the rotation around N-Cα-Xβ-Xγ • chi22) is the rotation around Cα-Xβ-Xγ-Xδ • chi33) is the rotation around Xβ-Xγ-Xδ-Xε Answer phi (Φ) = b, psi (Ψ) = a, chi11) = c, chi22) = d, and chi33) = e Question $19$ This question addresses the Biomolecular Visualization Framework theme(s) Atomic Geometry (AG), Topology and Connectivity (TC) Figure $12$ below shows an interactive iCn3D model of the atoms within 4A of the disulfide bond in Carbonic anhydrase from Neisseria gonorrhea (1KOQ). Rotate the model to determine the approximate chi33) dihedral angle. Hint: site down the S-S bond. Figure $12$: Atoms within 4A of the disulfide bond in Carbonic anhydrase from Neisseria gonorrhea (1KOQ). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...nutjP6EL2PubRA Answer The visually estimated chi33) angle for rotation around the S-S bond is 90o. Here is the actual angle (image made with Pymol) We mentioned previously that variants with higher thermal stability are likely to be more rigid and less flexible. Flexibility can be determined through analysis of molecular dynamic simulations and also by examining of the B factor values in PDB file. This number is a measure of the displacement of an atom from a mean The numbers in the last column in the file are called the temperature factors or B-factor. The B-factor describes the mean-square displacement, a measure of the displacement of an atom from an average value. If the atoms are more flexible, the electron density determined in x-ray structures is lower than if the atoms are more fixed, which gives high electron density. To make stabilizing disulfide bonds, investigators found site chains close enough that when mutated to cysteines could potentially form disulfide bonds. In addition, they search for such residues in surface loops (without alpha and beta structure) which are inherently more flexible. Introducing disulfide bonds into the loop would stabilize it and make it more rigid. Table $2$ below shows a description of the double cysteine CA variants in the study. Variant designation Position Wild-type residues Loop length Sum of B-factors T133C/D197C 133, 197 Thr/Asp 63 87.60 P56C/P156C 56, 156 Pro/Pro 99 80.82 N63C/P145C 63, 145 Asn/Pro 81 77.17 Table $2$: Description of the double cysteine CA variants in ngCA The locations of the side chair targeted for mutations to cysteine pairs are shown in Figure $13$ below. Figure $13$: 3D structure of ngCA and location of residue pairs for disulfide engineering The zinc (not shown)-coordinating histidine residues in the catalytic active site are shown in green. The proton shuttle histidine residue is shown in magenta. The native disulfide bond is colored yellow. An interactive iCn3D model of carbonic anhydrase from Neisseria gonorrhea (1KOQ) highlighting the 3 pairs of sidechains for mutations is shown in Figure $14$ below. Figure $14$: Carbonic anhydrase (Neisseria gonorrhea) with 3 paired side chains for engineered disulfide (1KOQ) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...KRP2tFJ1pV1FF6 The mutations were made and the wild-type proteins and three mutants were subjected to SDS-polyacrylamide gel electrophoresis (SDS-PAGE). The stained gels are shown in Figure $15$ below. Figure $15$: Expression and purification of disulfide CA variants. Panel (a) shows the expression of the protein in transformed cells 25 °C after IPTG induction and fractionated into soluble and insoluble fractions. SHuffle strain, an engineered E. coli strain that promotes cytoplasmic disulfide bond formation, was used. Panel (b) shows purification results. Each lane was loaded with 4 μg of each purified CA variant. The proteins were visualized with Coomassie blue staining after SDS-PAGE. The arrow indicates the position of the bands corresponding to ngCA variants. Lane: M, molecular weight marker; S, soluble fraction; IS, insoluble fraction. Question $20$ a. Interpret the results of the PAGE gels in Figure $15$ above b. How pure were the proteins based on the PAGE gel result in Panel (b). Can you infer from the gel that the proteins folded correctly? Answer a. It appears to show that a small fraction of the wild type and each mutant, especially the N63C/P145C pair, was found in the insoluble fraction. This might result from improper folding of the proteins in E. Coli, leading to hydrophobic side chain surface exposure and aggregation into insoluble "inclusion bodies". This appears to be just a minor issue. b. All the proteins appear very pure with some very small levels of contamination in the N63C/P145C. The PAGE results show the protein all have the same molecular weight but whether they folded to a native state with activity can not be determined. Nor can it be determined if the disulfide pairs in the mutants were made are if they were, were correctly paired. The investigators next determined if the expressed and purified wild-type and mutant proteins had the correct number of disulfide bonds. They did this by reacting the proteins in the absence and presence of dithiothreitol with DTNB or 5,5'-dithiobis(2-nitrobenzoic acid), also called Ellman's Reagent. Both structures are shown in Figure $16$ below. Figure $16$: Structures of DTT and DTNB DTT is a reducing agent that cleaves disulfide. Question $21$ Draw a mechanism showing the reaction of a disulfide with DTT. Answer Only surface and not buried free cysteines will be labeled unless the protein is unfolded to expose all the cysteines. DTNB reacts with free sulfhydryls to form the 2-nitro-5-thiobenzoic acid anion leaving group that absorbs at 412 nm. Question $22$ Draw a mechanism showing the reaction of free sulfhydryl (like Cys) with Ellman's reagent Answer Only surface and not buried free cysteines will be labeled unless the protein is unfolded to expose all the cysteines. The results of the reaction of the proteins with Ellmans's agent, in the presence and absence of DTT, are shown in Table $3$ below. CA variant Free thiol/protein (mol/mol) a Deduced no. S-S bonds −DTT +DTT Wild-type 0.06 ± 0.02 1.80 ± 0.14 ? T133C/D197C 0.08 ± 0.02 3.79 ± 0.22 ? P56C/P156C 0.06 ± 0.03 3.89 ± 0.06 ? N63C/P145C 0.08 ± 0.03 3.75 ± 0.05 ? Table $3$: Analysis of disulfides in CA using Ellman's reagent. aNumbers are represented in mean ± SD. Question $23$ How many S-S would you deduce from the table are present in the wild-type and mutant enzymes? Did the correct disulfide bonds form? Explain your answers Answer DTT reduces the disulfide in protein. For each disulfide, two free Cys side chains are made. The molar ratio of CysSH/CA for the wild-type is 1.8 in the presence of DTT. The value is very close to the expected value of 2. For the mutants, the ratio is about 3.8 in the presence of DTT, suggesting 4 free Cys consistent with 2 disulfide bonds. Table $4$ below shows the catalytic activities of the disulfide CA variants at 25 °C. CA variant CO2 hydration activity Relative esterase activity a kcat × 10−4 (s−1) K M (mM) kcat /KM × 10−6(M−1 s−1) Wild-type 1.00 1.44 14.2 1.01 T133C/D197C 1.49 1.97 16.7 1.18 P56C/P156C 1.03 1.44 16.9 0.85 N63C/P145C 0.55 0.27 17.3 0.16 Table $4$: catalytic activities of the disulfide CA variants at 25 °C aThe specific activity of the wild-type corresponds to 0.22 U/μmol-enzyme. Question $24$ Why did the investigators conduct this experiment? Interpret the results Answer All of the previous results suggest that the mutant proteins were made and had the correct number of double bonds, but the experiments could not tell if the bond pairs were correct. For example, did the T133C/D197C contain a native (C28-C181) and mutant (C133-C197) bond and not another combination? Activity is an excellent predictor of structure. All but one mutant retained nominal activity, as evidenced by a comparison of the rat constants. The N63C/P145C had a 6x lower kcat, but even then it is close to diffusion-controlled. Now comes the big question: were the investigators able to engineer thermal stability into the carbonic anhydrase? Experimental results to show the thermostability of the disulfide CA mutants are shown in Figure $17$ below. Figure $17$: Thermostability of the disulfide CA variants Panel (a) shows short-term kinetic stability. The enzyme solutions (40 μM) were incubated for 30 min at different temperatures, and the residual activities were measured by esterase activity assay. Activities of 100% correspond to untreated samples. Panel (b) shows long-term kinetic stability at 70 °C. The half-lives (t1/2) of the CA variants were estimated by fitting the experimental data to an exponential decay curve. Each value represents the mean of at least three independent experiments, and the error bars represent the standard deviations. Question $25$ Analyze the results in Panels (a) an (b). Which protein was most thermostable over the short (30 minute) and long (hours) incubating time at elevated temperatures? Answer At 80 °C, all the mutants showed increases thermostability to short term (30 minute) heating, but one, N63C/P145C was exceptionally thermostable. Longer time courses for heating at 70°C showed that the N63C/P145C was again far more stable over time. Its t1/2 was 31.4 h, compared to the values between 4-6 h for the others. Panel (c) shows heat-induced denaturation of disulfide CA variants. Temperature-dependent changes of the circular dichroism ellipticity were recorded at 220 nm on CD spectrometer. The denaturation curves were normalized to the fraction of unfolded protein. The horizontal dashed line indicates the point at which the fraction of unfolded protein is 0.5. The vertical dashed lines point to TM values. Panel (d) shows the overall RMSD of disulfide variants from molecular dynamic simulations performed at 400 K for 20 ns. Question $26$ Analyze the results in Panel (c). What do changes in the CD helicity show? Which protein was most thermostable based on TM values? Is the decrease in enzyme activity in panels (a) and (b) result from the denaturation of the protein? Answer CD measurements can give a measure of the retention of secondary structure (alpha helices and beta structure) on denaturation. The CD spectrum for different secondary structures is shown below (From Chapter 3.5). The curves were normalized to fit on a 0-1 scale on the y axis, which then gives a measure of percent denaturation. The temperature half-way up is the TM, or "melting temperature, at which an equilibrium mixture would contain half native and half denatured protein (true for a small protein with no intermediates). The TM values were for the wild-type, T133C/D197C, P56C/P156C, and N63C/P145C mutants 73.6 °C 74.7 °C, 77.4 °C, and 81.4 °C. These parallel the t1/2 values for enzyme activities, and support the idea that denaturation led to inactivation of the enzyme. Question $27$ Analyze the molecular dynamics simulation results in Panel (d) Answer The molecular dynamic simulations for all the proteins soon reach equilibrium values as indicated in the plateaus of average room mean square deviation of the protein backbone. The overall molecular root-mean-square deviation (RMSD) of N63C/P145C was the lowest, indicating that it was most rigid. This is in accord with the idea that increased flexibility destabilizes a protein and engineering a disulfide into makes it more rigid and hence more stable to temperature increases. The results are in accordance with the other experiments that show the N63C/P145C was the most thermostable. " In addition, T133C/D197C showed the highest values in both the overall and the residual RMSD (Fig 3D). This may explain and correlate with the increased activity of T133C/D197C (Ta) and the increased ΔS of unfolding" You may remember from both introductory chemistry and from Chapter 4.12, that you can calculate the thermodynamic parameters, ΔHo and ΔSo for N ↔D at room temperature from thermal denaturation curves using the van 't Hoff equation. In this case, Keq values can be calculated from thermal denaturation curves by monitoring change in CD signal at 220 nm, and applying this equation (also from Chapter 3.12). K_{e q}=\frac{[D]_{e q}}{[N]_{e q}}=\frac{f_D}{f_N}=\frac{f_D}{1-f_D} From this, we can calculate ΔG0. \Delta \mathrm{G}^0=-\mathrm{R} \operatorname{Tln} \mathrm{K}_{\mathrm{eq}}=-\mathrm{R} \operatorname{Tln}\left[\frac{\mathrm{f}_{\mathrm{D}}}{1-\mathrm{f}_{\mathrm{D}}}\right] Knowing Keq, ΔH0, DS0 can be calculated as shown below. A semi-log plot of lnKeq vs 1/T is a straight line with a slope of - ΔH0R and a y-intercept of + ΔS0/R, where R is the ideal gas constant. \begin{gathered} \Delta \mathrm{G}^{0}=\Delta \mathrm{H}^{0}-\mathrm{T} \Delta \mathrm{S}^{0}=-\mathrm{RTln} \mathrm{K}_{\mathrm{eq}} \ \ln \mathrm{K}_{\mathrm{eq}}=-\frac{\Delta \mathrm{H}^{0}-\mathrm{T} \Delta \mathrm{S}^{0}}{\mathrm{RT}} \ \ln \mathrm{K}_{\mathrm{eq}}=-\frac{\Delta \mathrm{H}^{0}}{\mathrm{RT}}+\frac{\Delta \mathrm{S}^{0}}{\mathrm{R}} \end{gathered} The equation below shows that the derivative of equation (8) with respect to 1/T (i.e. the slope of equation 8 plotted as lnKeq vs 1/T) is indeed -ΔH0/R. Equation (9) is the van 't Hoff equation, and the calculated value of the enthalpy change is termed the van 't Hoff enthalpy, ΔH0vHoff. \frac{d \ln \mathrm{K}_{\mathrm{eq}}}{d(1 / \mathrm{T})}=-\frac{\Delta \mathrm{H}^{0}}{\mathrm{R}}=-\frac{\Delta \mathrm{H}_{\mathrm{vHoff}}^{0}}{\mathrm{R}} Using this method, the thermodynamic parameters for unfolding of the protein were calculated. The results are shown in Table $4$ below. CA variant Melting temperature, TM (°C) Enthalpy change of unfolding, ΔH (kcal mol−1) Entropy change of unfolding, ΔS (kcal mol−1 K−1) Wild-type 73.6 48.8 0.141 T133C/D197C 74.7 52.8 0.153 P56C/P156C 77.4 35.1 0.091 N63C/P145C 81.4 30.0 0.085 Table $4$: Thermodynamic parameters for protein unfolding for WT and mutant CAs Question $28$ Which effects, enthalpy or entropy of unfolding, were associated with the increased thermal stability of the mutants compared to the wild-type protein. Remember were are considering the denaturation reaction, N↔ D. Answer For the reaction N ↔ D, the ΔH0 values were all positive, indicating the enthalpy changes favored the native state, not the denatured state. In contrast, the other two mutants were enthalpically destabilized compared to the wild-type as their ΔH0 were less positive so compared to the wild-type. The prime stabilizer of the native state was the lower entropy (hence a less negative and favored -TΔS0 for the denaturation reaction. This makes sense in these mutants are more rigid and would experience less loss of "conformational entropy). P56C/P156C and N63C/P145C exhibited lower ΔH (destabilizing) and ΔS (stabilizing), showing that the decreased entropic change of unfolding (i.e., the loss of conformational entropy of the unfolded state) by the disulfide bridge was the primary factor for the thermostabilization. These results are not surprising because design strategies aiming ‘entropic stabilization’ such as disulfide engineering do not always result in engineered proteins ideally with lower ΔS and unchanged ΔH. These results are in accord with the observation that N63C/P145C was the most thermostable variant and that T133C/D197C showed the highest values in both the overall and the residual RMSD (Fig. 3d). This may explain and correlate with the increased activity of T133C/D197C and the increased ΔS of unfolding. Finally, the enzymatic activity of the wild-type and mutants CAs (using a small ester substrate) were studied as a function of temperature. The relative activity of the wild-type and all 3 disulfide mutants are plotted as a function of temperature in the histogram graphs shown in Figure $18$ below. Figure $18$: Effect of temperature on the activity of disulfide CA variants. Esterase activities of disulfide variants were measured at each temperature and normalized to the activity of each enzyme at 25 °C. Each value represents the mean of three independent experiments, and the error bars represent the standard deviations. If you plotted the data as curves, you would get bell-shaped graphs. Question $29$ Explain why the histogram plots (and line plots if they were drawn) are bell-shaped. Are the results in accordance with the previous results. Answer Yes. Most chemical reaction show an increase in rate with increasing temperatures until competing reactions take precedence. For an enzyme-catalyzed reaction, that competing reaction is denaturation, which decreases the rate. Yes the graphs are in accord with the previous results. The N63C/P145C certainly stands out as the best mutant. The authors write that "considering the shifted optimal temperature and the thermoactivation as well as the enhanced thermostability, the disulfide engineered α-type CA with Cys63-Cys145 can be a promising biocatalyst for efficient CO2 sequestration performed under high temperature conditions." Disulfide engineering Craig, D.B., Dombkowski, A.A. Disulfide by Design 2.0: a web-based tool for disulfide engineering in proteins. BMC Bioinformatics 14, 346 (2013). https://doi.org/10.1186/1471-2105-14-346. Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), Several computational programs have been developed to determine amino acid pairs that could be mutated to high-temperature stabilizing disulfide bonds. The stabilizing effects appear largest when the disulfide bond is made within the largest (and most flexible) loops (between 25-75 residues). These loops also had the highest residue B-factors. In selecting pairs to form engineered disulfide, not only proximity (distance) but also geometry (torsion angles) of the resulting disulfide bond are important. We saw this previously in the analysis of the energy of butane rotamers, as illustrated in Figure $19$ below. Figure $19$: Newman projections for butane Programs to determine amino acid pairs to mutate for disulfide bond formation test S-S bond torsional stability by determining the torsion angle χ3 for the S-S bond. Evaluation of a database of many native proteins shows the χ3 angle are centered in two major peaks at -87 and +97 degrees, as shown in Figure $20$ below Figure $30$: Distribution of χ 3 torsion angles observed in 1505 native disulfide bonds found in 331 PDB protein structures. Peaks occur at -87 and +97 degrees. Craig, D.B., Dombkowski, A.A. Disulfide by Design 2.0: a web-based tool for disulfide engineering in proteins. BMC Bioinformatics 14, 346 (2013). https://doi.org/10.1186/1471-2105-14-346. Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), Question $4$ Does your calculated value of χ3 for the native disulfide in carbonic anhydrase from Neisseria gonorrhea follow the observed angles? Answer At about 90o so yes it does. An empirical energy equation was developed to determine the E vs χ3 dihedral angle for disulfide bonds (Craig, D.B., Dombkowski, A.A., ibid). It is shown below. E\left(\chi_3\right)=4.0\left[1-\cos \left(1.957\left[\chi_3+87\right]\right)\right] A graph of the equation made with Excel is shown in Figure $21$ below Figure $21$: E vs χ3 dihedral angle for disulfide bonds from empirical equation Figure $22$ shows the distribution of energy values in the 1505 native disulfide bonds using our updated function. The study by Craig and Dombkowski showed that almost all (90%) of disulfides in native proteins in the PDB have an energy < 2.2 kcal/mol, so this metric could be used to determine possible disulfide bond pairs created by mutagenesis. Figure $22$: Distribution of the disulfide bond energy calculated for 1505 native disulfide bonds in our survey set using the DbD2 energy function. The mean value is 1.0 kcal/mol, and the 90th percentile is 2.2 kcal/mol. (Craig, D.B., Dombkowski, A.A., ibid) Question $31$ You calculated the approximate value for χ3 dihedral for the C28-C181 native disulfide bond form in Carbonic anhydrase from Neisseria gonorrhea using iCn3D in Question x above. Determine the approximate energy from the empirical function graph in Figure x above for that χ3 dihedral. What percent of native disulfide bonds have that particular calculated energy in the 1505 native disulfide surveyed? Answer The estimated χ3 dihedral was 90o. The energy for that χ3 angle would be approximately <0.1kcal/mol, which reflects the energies of about 150/1505 or 10% of the disulfides in the database. It is hence in the most stable range of disulfides, based only on the χ3 dihedral angle. Yet there are other important parameters as well that would affect the energy of the disulfide bond in the protein. Figure $23$ below shows a comparison of native residue B-factors in stabilizing and destabilizing engineered disulfide bonds Figure $23$: Comparison of native residue B-factors in stabilizing and destabilizing engineered disulfide bonds. The native structures associated with engineered disulfides previously reported as stabilizing (S) or destabilizing (D), based on experimental evidence, were analyzed with DbD2. The mean B-factor for residues involved in stabilizing disulfide bonds was 31.6 compared with 16.5 for those involved in destabilizing bonds, P = 0.066. (Craig, D.B., Dombkowski, A.A., ibid) Question $32$ Which proteins were more stabilized by engineered disulfide, those with higher or low B factors. Answer Proteins with engineered disulfides that increase stability have higher B-factors. This makes sense in that proteins with higher B-factors and hence mobility would be predicted to alter conformation and potentially denature more readily. Another Paper: Prediction of disulfide bond engineering sites using a machine learning method https://www.nature.com/articles/s41598-020-67230-z Gao, X., Dong, X., Li, X. et al. Prediction of disulfide bond engineering sites using a machine learning method. Sci Rep 10, 10330 (2020). https://doi.org/10.1038/s41598-020-67230-z. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ The amount of data present in a single PDB file is very large, but it is nothing compared to the collective data in all PDB files. If only we could extract empirical rules from the collective PDB files that govern disulfide bond formation. It turns out we can with machine learning and artificial intelligence that can be used to develop and train predictive algorithms. Machine learning has been used to predict amino acid pairs for cysteine mutations to form engineered disulfide bonds. It recognizes 99% of natural disulfide bonds. residues. It uses these parameters: • distances between the alpha-carbons and the beta-carbons of the bonded cysteine residues • three torsion angles around the disulfide bonds (χ1ss1’). An example of one variable that helps define the stability of disulfide bonds is the distances between the Cα atoms for the disulfide-bonded cysteines, as shown in Figure $24$ below. Figure $24$: The histogram of distances between Cα atoms of disulfide-bonded cysteines. The distances range from 3.0 Å and 7.5 Å. Knowing this would constrain the number of choices for paired amino acid side chains for mutagenesis to produce disulfide. Machine learning can also be used to find other distance constraints to optimize mutagensis experiments. Figure $25$ below shows a graph of 10 different distances and their relative importance in determining disulfide bond stability. Figure $25$: The relevance of the distance features to the classification outcome. Out of the 45 unique distances, 20 distances have negligible influence on the classification performance. The distances between Cβ and main-chain atoms of the pairing residue are important features in disulfide bond classifications. An interactive iCn3D model of carbonic anhydrase from Neisseria gonorrhea (1KOQ) highlighting two pairs of amino acids identified by machine learning as candidates for mutations to disulfide-bonded cysteines is shown in Figure $26$ below. Figure $26$: Carbonic anhydrase from Neisseria gonorrhea (1KOQ) highlighting two pairs of amino acids identified by machine learning as candidates for mutations to disulfide-bonded cysteines (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...ZH6Tv1urwkKhz8 Question $33$ This question addresses the Biomolecular Visualization Framework theme(s) Atomic Geometry (AG), Macromolecular Building Blocks (MB) Using the data in Figure $25$ and measurements made using the iCn3D model above (Figure $25$) , determine which pair would be most likely engineered into a disulfide bond. Complete the table below with the distances you made using iCn3D. iCn3D instructions: Open the external link and follow these instructions Trackpad and Mouse Controls rotate: click and drag (mouse: left click and drag) zoom: pinch and spread (mouse: rotate the scroll wheel) translate: two-finger click and drag (mouse: right click and drag) Re-center: left click View from the top menu bar, then select “Center Selection” •Note: ctrl-click on a PC = command-click on Mac; alt-click on PC = option click on Mac Instructions 1. Zoom to clearly see the amino acid pair for distance measures 2. From the top menu bar, choose Analysis, Distance, distance between 2 atoms 3. Pick the appropriate 2 atoms for measure distance by holding down the Alt key and selecting both 4. Record the distances in the table below. Mutation Pair CB1-CA2 (Å) CB1-CB2 (Å) CA1-CB2 (Å) 1 (L137) - 2 (W141) 1 (Y54) - 1 (S160) Answer Mutation Pair CB1-CA2 (Å) CB1-CB2 (Å) CA1-CB2 (Å) 1 (L137) - 2 (W141) 6 5.2 5.3 1 (Y54) - 1 (S160) 4.1 4.1 5.0 Errors, Misformation and Disinformation Under construction An interactive iCn3D model of the anti-arsonate germline antibody 36-65 in complex with a phage display derived dodecapeptide KLASIPTHTSPL without added hydrogens (2A6I) Figure \(5\): Anti-arsonate germline antibody 36-65 in complex with a phage display derived dodecapeptide KLASIPTHTSPL without added hydrogens (2A6I). (Copyright; author via source).  Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...hi5pLnMmq6MEJ6 Download this file and open in iCn3D (File, Open File, iCn3D PNG Image) to see a model with attached hydrogen atoms 2.5A resolution, disallowed regions 1% Link to interpret quality me, districs Figure \(5\) is the Percentile scores (ranging between 0-100) for global validation metrics of the entry a Figure \(5\): geometric issues observed across the polymeric chains and their fit to the electron density. https://www.rcsb.org/structure/2a6i Rfree is a measure of the quality of a model from X-ray crystallographic data The Figure \(5\)below summarizes the geometric issues observed across the polymeric chains and their fit to the electron density. The red, orange, yellow, and green segments on the lower bar indicate the fraction of residues that contain outliers for > 3, 2, 1 and 0 types of geometric quality criteria respectively. A grey segment represents the fraction of residues that are not modeled. The numeric value for each fraction is indicated below the corresponding segment, with a dot representing fractions <5% The upper red bar (where present) indicates the fraction of residues that have poor fitt to the electron density. The numeric value is given above the bar Figure \(5\)"  https://www.rcsb.org/structure/2a6i KLASIPTHTSPL only 9 observable electron density (S4 to end) updated version:  5VGA Exercise \(1\) Show van der Waals steric clashes in the protein using the program Jsmol available at this link • check the with hydrogens box on the right-hand side • click in the load mmCIF by PDB ID and input the 2a6i (small letters) • select the Clashes button on the left • rotate the image to display the greatest density of clashes to the right. • Select the PNG + Jmol  button on the right-hand side to download an image showing the clashes • hover over the region with the greatest number of clashes to identify amino acids in this region.  Which chain (A, B, P) is involved in the most clashes? Answer The right-hand side with the greatest density of clashes shows the source of most clashes is the bound peptide P.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/Global_Challenges_-_Literature-Based_Guided_Assessments_(LGAs)/2.__Global_Challenges_-_Climate_Change%3A__Carbo.txt

Under Construction Glutamatergic transmission has been implicated in the pathophysiology of PTSD, particularly in the effects of N-methyl-D-aspartate receptor (NMDAR) signaling on the synaptic plasticity underlying learning and memory [13]. NMDARs comprise two GluN1 subunits and two GluN2 (A-D) or GluN3 (A, B) subunits. In adult forebrain regions, GluN2A and GluN2B are the main subunits forming receptor complexes with GluN1 at excitatory synapses. GluN2B-containing NMDARs play a preferential role in inducing synaptic plasticity, which is critical for the extinction of fear memories [1415]. Systemic injection of GluN2B-specific NMDAR antagonists ((RS)-3-(2-carboxypiperazin-4-yl)-propyl-1-phosphonic acid, ifenprodil) can impair the retention of fear extinction learning. GluN2B-containing NMDARs in both the amygdala and medial prefrontal cortex (mPFC) are also involved in reducing fear during extinction, whereas GluN2A-containing NMDARs play a greater role in the initial formation and/or stabilization of learned fear [15]. Rodent studies demonstrate that GluN2B subunit-containing NMDARs play pivotal roles in fear extinction learning. https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4263351/"tetrameric complexes mainly composed of NMDA receptor GluNR1 and GluNR2 subunits with the NR2 subunits modifying the activity of the receptor. " N-methyl-d-aspartate (NMDA) receptors are Hebbian-like coincidence detectors, requiring binding of glycine and glutamate in combination with the relief of voltage-dependent magnesium block to open an ion conductive pore across the membrane bilayer. NMDA receptors are Hebbian-like coincidence detectors, requiring the binding of glycine and glutamate to GluN1 and GluN2 subunits, respectively, combined with membrane depolarization to relieve magnesium block . Activation of the receptor opens a cation-selective, calcium permeable channel, thus causing further depolarization of the cell membrane and influx of calcium . NMDA receptors are obligatory heterotetrameric assemblies, typically composed of two glycine-binding GluN1 subunits and two glutamate-binding GluN2A-D subunits, with the GluN1/GluN2A/GluN2B complex the predominate receptor at hippocampal synapses. Glycine-and d-serine-binding GluN3 subunits are additional subunits, expressed throughout the nervous system but with roles less well defined in comparison to the GluN1/GluN2 assemblies. A hallmark of NMDA receptors, by contrast with AMPA and kainate receptors, is a wide spectrum of allosteric modulation, from nanomolar concentrations of zinc, to the small molecule ifenprodil, polyamines and protons and to voltage-dependent ion channel block by MK-801, ketamine and memantine. The coordinates and structure factors for the structure have been deposited in the Protein Data Bank under accession code 4TLL and 4TLM for Structure 1 and Structure 2, respectively. These receptors have two GluN1 subunits and two GluN2 (A-D) or GluN3 (A, B) subunits.  In the forebrain, GluN2A and B form complex with GluN1 at synapses, with the B subunit playing a role in synaptic plasticity.  Synaptic plasticity is necessary to remove "hard-wired" fear circuits. A goal of PTSD therapies is the extinction of the previously acquired feared memories through learning.  Learning, and more specifically extinction, requires synaptic plasticity. If one goal of PTSD treatment is the extinction of fear memories, then drugs that target GluN2B are potentially useful.  Antagonist (such asd ifenprodil) of GluN2B seem to decrease the ability to extinguish fear retention memories.  In other words, the learning and synaptic plasticity need to attenuate the fear memories are inhibited by the antagonist.  GluN2B in the NMDAR receptors in the amygdala and medial prefrontal cortex appear to be involved in reducing feat during extinction.  IN contact, GluN2A seem to be an important role in forming and stabilizing the learned fear response. "Glutamatergic transmission has been implicated in the pathophysiology of PTSD, particularly in the effects of N-methyl-D-aspartate receptor (NMDAR) signaling on the synaptic plasticity underlying learning and memory [13]. NMDARs comprise two GluN1 subunits and two GluN2 (A-D) or GluN3 (A, B) subunits. In adult forebrain regions, GluN2A and GluN2B are the main subunits forming receptor complexes with GluN1 at excitatory synapses. GluN2B-containing NMDARs play a preferential role in inducing synaptic plasticity, which is critical for the extinction of fear memories [1415]. Systemic injection of GluN2B-specific NMDAR antagonists ((RS)-3-(2-carboxypiperazin-4-yl)-propyl-1-phosphonic acid, ifenprodil) can impair the retention of fear extinction learning. GluN2B-containing NMDARs in both the amygdala and medial prefrontal cortex (mPFC) are also involved in reducing fear during extinction, whereas GluN2A-containing NMDARs play a greater role in the initial formation and/or stabilization of learned fear [15]. Rodent studies demonstrate that GluN2B subunit-containing NMDARs play pivotal roles in fear extinction learning." Describe how the NMDA receptor functions, and how it implements the Hebbian model of learning at the synaptic level. http://charlesfrye.github.io/Foundat...roscience//29/ https://www.pnas.org/doi/full/10.1073/pnas.95.12.7145 https://pubmed.ncbi.nlm.nih.gov/15888440/ https://www.ncbi.nlm.nih.gov/pmc/articles/PMC4263351/ Ras Questions 1. Where is Ras located in the cell?  What causes Ras to be localized here? 2. How does Ras get activated? 3. What are the three dimensional differences between GDP and GTP bound Ras.  Why is the GTP bound form “active?” 4. What does Ras bound to GTP bind to?  What are the net effects of this in terms of signal transduction? 5. What reaction does Ras catalyze as an enzyme? 6. What type of reaction is a GAP catalyzing? GEF? a. Are they opposite reactions? What is different (enzymatic control) 7. GAP is an acronym for GTPase activating protein. What GTPase is being activated? 8. Would RAS “turn off” without a GAP present? Why is this critical? 9. What molecule (GTP or GDP) would be bound to RAS if there were no GAPs or GEFs present? Why? 10. Why does a GAP increase the enzymatic activity RAS?  What does a GAP provide that aids in the chemistry of the reaction? Paper about targeting phosphorylation and cancer therapy:  https://www.ncbi.nlm.nih.gov/pmc/articles/PMC8642438/ 1. Ras can be phosphorylated at Tyr 32 and Tyr64 by Src kinase, which leads to inhibition of binding to Raf and increased GTP hydrolysis.  Provide a rationale for why this post-translational modification plays a role in the overall function of Ras.  (Not sure what to ask here- there seems to be a number of kinases involved that do different things as explained in the paper above) 2.  There are mutations in Ras, including Gly 12, Gly13, and Gln 61 which impair GTPase activity and GAP-mediated GTP hydrolysis.  Predict what changes this would cause in the cell. 3. Think about designing a small molecule drug that affects the Ras signaling pathway and treats cancer.  What proteins could you target?  For each protein target,  should the drug increase or decrease the activity of its target?  Explain your answers. Fig 1 Fig 2 Fiog 3

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/Signal_Transduction_Problems/NMDA_Receptor_-_Under_Construction.txt

Disulfide engineering The first set of questions below are based on this reference as noted: Craig, D.B., Dombkowski, A.A. Disulfide by Design 2.0: a web-based tool for disulfide engineering in proteins. BMC Bioinformatics 14, 346 (2013). https://doi.org/10.1186/1471-2105-14-346. Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0) Several computational programs have been developed to determine amino acid pairs that could be mutated to high-temperature stabilizing disulfide bonds. The stabilizing effects appear largest when the disulfide bond is made within the largest (and most flexible) loops (between 25-75 residues). These loops also had the highest residue B-factors. In selecting pairs to form engineered disulfide, not only proximity (distance) but also geometry (torsion angles) of the resulting disulfide bond are important. We saw this previously in the energy analysis of butane rotamers, as illustrated in Figure $1$ below. Figure $1$: Newman projections for butane Programs to determine amino acid pairs to mutate for disulfide bond formation test S-S bond torsional stability by determining the torsion angle χ3 for the S-S bond. Evaluation of a database of many native proteins shows the χ3 angle are centered in two major peaks at -87 and +97 degrees, as shown in Figure $2$ below Figure $2$: Distribution of χ 3 torsion angles observed in 1505 native disulfide bonds found in 331 PDB protein structures. Peaks occur at -87 and +97 degrees. Craig, D.B., Dombkowski, A.A. Disulfide by Design 2.0: a web-based tool for disulfide engineering in proteins. BMC Bioinformatics 14, 346 (2013). https://doi.org/10.1186/1471-2105-14-346. Creative Commons Attribution License (http://creativecommons.org/licenses/by/2.0), Question $1$ Does your calculated value of χ3 for the native disulfide in carbonic anhydrase from Neisseria gonorrhea follow the observed angles? Answer At about 90o so yes it does. An empirical energy equation was developed to determine the E vs χ3 dihedral angle for disulfide bonds. The equation is shown below (Craig, D.B., Dombkowski, A.A. ibid.). E\left(\chi_3\right)=4.0\left[1-\cos \left(1.957\left[\chi_3+87\right]\right)\right] A graph of the equation made with Excel is shown in Figure $3$ below Figure $3$: E vs χ3 dihedral angle for disulfide bonds from empirical equation Figure $4$ below shows the distribution of energy values in the 1505 native disulfide bonds using our updated function. The study by Craig and Dombkowski showed that almost all (90%) of disulfides in native proteins in the PDB have an energy < 2.2 kcal/mol, so this metric could be used to determine possible disulfide bond pairs created by mutagenesis. Figure $4$: Distribution of the disulfide bond energy calculated for 1505 native disulfide bonds. The mean value is 1.0 kcal/mol, and the 90th percentile is 2.2 kcal/mol. Craig, D.B., Dombkowski, A.A. ibid. Question $2$ You calculated the approximate value for χ3 dihedral for the C28-C181 native disulfide bond form in Carbonic anhydrase from Neisseria gonorrhea using iCn3D in Question x above. Determine the approximate energy from the empirical function graph in Figure x above for that χ3 dihedral. What percent of native disulfide bonds have that particular calculated energy in the 1505 native disulfide surveyed? Answer The estimated χ3 dihedral was 90o. The energy for that χ3 angle would be approximately <0.1kcal/mol, which reflects the energies of about 150/1505 or 10% of the disulfides in the database. It is hence in the most stable range of disulfides, based only on the χ3 dihedral angle. Yet there are other important parameters as well that would affect the energy of the disulfide bond in the protein. Figure $5$ below shows a comparison of native residue B-factors in stabilizing and destabilizing engineered disulfide bonds Figure $5$: Comparison of native residue B-factors in stabilizing and destabilizing engineered disulfide bonds. Analysis of previously reported proteins with stabilizing (S) or destabilizing (D) engineered disulfide bonds. The mean B-factor for residues in proteins with stabilizing disulfide bonds was 31.6 compared with 16.5 for those involved in destabilizing bonds. Craig, D.B., Dombkowski, A.A. ibid. Question $3$ Which proteins were more stabilized by engineered disulfide, those with higher or low B factors. Answer Proteins with engineered disulfides that increase stability have higher B-factors. This makes sense in that proteins with higher B-factors and hence mobility would be predicted to alter conformation and potentially denature more readily. Prediction of disulfide bond engineering sites using a machine learning method This set of questions based on this reference: Gao, X., Dong, X., Li, X. et al. Prediction of disulfide bond engineering sites using a machine learning method. Sci Rep 10, 10330 (2020). https://doi.org/10.1038/s41598-020-67230-z. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ The amount of data present in a single PDB file is very large, but it is nothing compared to the collective data in all PDB files. If only we could extract empirical rules from the collective PDB files that govern disulfide bond formation. It turns out we can with machine learning and artificial intelligence that can be used to develop and train predictive algorithms. Machine learning has been used to predict amino acid pairs for cysteine mutations to form engineered disulfide bonds. It recognizes 99% of natural disulfide bonds. residues. It uses these parameters: • distances between the alpha-carbons and the beta-carbons of the bonded cysteine residues • three torsion angles around the disulfide bonds (χ1ss1’). An example of one variable that helps define the stability of disulfide bonds is the distances between the Cα atoms for the disulfide-bonded cysteines, as shown in Figure $6$ below. Figure $6$: The histogram of distances between Cα atoms of disulfide-bonded cysteines. Gao, X., Dong, X., Li, X. et al. Prediction of disulfide bond engineering sites using a machine learning method. Sci Rep 10, 10330 (2020). https://doi.org/10.1038/s41598-020-67230-z. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ The distances range from 3.0 Å and 7.5 Å. Knowing this would constrain the number of choices for paired amino acid side chains for mutagenesis to produce disulfide. Machine learning can also be used to find other distance constraints to optimize mutagensis experiments. Figure $7$ below shows a graph of 10 different distances and their relative importance in determining disulfide bond stability. An interactive iCn3D model of carbonic anhydrase from Neisseria gonorrhea (1KOQ) highlighting two pairs of amino acids identified by machine learning as candidates for mutations to disulfide-bonded cysteines is shown in Figure $8$ below. Figure $8$: Carbonic anhydrase from Neisseria gonorrhea (1KOQ) highlighting two pairs of amino acids identified by machine learning as candidates for mutations to disulfide-bonded cysteines (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...ZH6Tv1urwkKhz8 Question $4$ This question addresses the Biomolecular Visualization Framework theme(s) Atomic Geometry (AG), Macromolecular Building Blocks (MB) Using the data in Figure $25$ and measurements made using the iCn3D model above (Figure $25$) , determine which pair would be most likely engineered into a disulfide bond. Complete the table below with the distances you made using iCn3D. iCn3D instructions: Open the external link and follow these instructions Trackpad and Mouse Controls rotate: click and drag (mouse: left click and drag) zoom: pinch and spread (mouse: rotate the scroll wheel) translate: two-finger click and drag (mouse: right click and drag) Re-center: left click View from the top menu bar, then select “Center Selection” •Note: ctrl-click on a PC = command-click on Mac; alt-click on PC = option click on Mac Instructions 1. Zoom to clearly see the amino acid pair for distance measures 2. From the top menu bar, choose Analysis, Distance, distance between 2 atoms 3. Pick the appropriate 2 atoms for measure distance by holding down the Alt key and selecting both 4. Record the distances in the table below. Mutation Pair CB1-CA2 (Å) CB1-CB2 (Å) CA1-CB2 (Å) 1 (L137) - 2 (W141) 1 (Y54) - 1 (S160) Answer Mutation Pair CB1-CA2 (Å) CB1-CB2 (Å) CA1-CB2 (Å) 1 (L137) - 2 (W141) 6 5.2 5.3 1 (Y54) - 1 (S160) 4.1 4.1 5.0

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/Structure%2F%2FFunction_-_Protein_Problems/Disulfide_Bonds.txt

Written by Henry Jakubowski, Emily Schmitt Lavin, Arthur Sikora, and Subhasish Chatterjee Introduction Eukaryotic voltage-gated sodium (NaV) channels generate and sustain action potentials in nerve and muscle cells by moving Na+ ions from the outside to the inside of the cell.  This increases and makes positive the transmembrane potential of the cell, which at rest is approximately -70 mV (more negative inside).  Once activated, the channel undergoes a fast inactivation (1-2 ms), without which the firing of nerves and muscles becomes dysregulated, a potentially lethal effect.  Please view the information in Chapter 11.3 on the voltage-gated sodium channel before you do these guided assessment activities. The questions below are derived from a paper from Jiang et al. on the structure and properties of the α-scorpion toxin LqhIII (MW 7,000) bound to rat cardiac sodium channel NaV1.5 (MW 227,000) by. (Jiang, D., Tonggu, L., Gamal El-Din, T.M. et al. Structural basis for voltage-sensor trapping of the cardiac sodium channel by a deathstalker scorpion toxin. Nat Commun 12, 128 (2021). https://doi.org/10.1038/s41467-020-20078-3.  Creative Commons Attribution 4.0 International License.  http://creativecommons.org/licenses/by/4.0/) The Lqh toxin is made by Leiurus quinquestriatus hebraeus. It is found in North Africa, the Middle East, and Western India and is shown below. The study shows how the deathstalker scorpion (LqhIII) toxin inhibits the fast inactivation of cardiac sodium channels (Nav1.5). In other words, you will see how the toxin keeps the channel open longer than it would be open in its absence. • In the absence of toxin, the sodium channel NaV1.5 returns in 1-2 ms to an inactive state when an 'inactivation gate" moves to occlude the open pore. • The α-scorpion toxin LqhIII inhibits the return of the channel to the inactive state.   Since the toxin inhibits the channel's fast inactivation of Na+ ion flow into the cell, the channel stays open longer. The toxin leads to the inhibition of the normal fast inhibition (inactivation) of the channel. Hence the channel stays open (activated) longer.  This is analogous to the statement that the enemy of my enemy is my friend! Techniques to study the Na Channel For the experiments described in the paper, the rat sodium cardiac channel NaV1.5 was purified, its structure determined, and its functional properties (the regulated movement of Na+ to the inside of the cell - electrophysiology) in human epithelial cells measured.   The protein is expressed in rat cardiac cells and is found in the cell membrane of the cell.  The iCn3D image below shows the alpha chain of rat cardiac NaV1.5 (6UZ3) embedded in a simple bilayer (DMPC) to model how it might appear in a cardiac cell membrane bilayer. PDB coordinates based on S. Jo, T. Kim, V.G. Iyer, and W. Im (2008). CHARMM-GUI: A Web-based Graphical User Interface for CHARMM. J. Comput. Chem. 29:1859-1865.  S. Jo, T. Kim, and W. Im (2007) Automated Builder and Database of Protein/Membrane Complexes for Molecular Dynamics Simulations. PLoS ONE 2(9):e880 Exercise \(1\) Complete the flow chart below to show two different approaches that could be used to purify the protein and prepare it for structural and functional studies.   On the left show steps you could use to directly purify the protein from rat hearts.  On the right use the rat gene (Scn5a) for the channel as the starting point for purification. Here are some links for review if needed. Answer From heart tissue: From heart tissue DNA Here are some review links: Exercise \(2\) Key components of the buffer solution used to purify the channel include HEPES and 1% (w/v) n-dodecyl-β-D-maltopyranoside.  Their structures are shown below. Describe the role of each. Answer . Exercise \(3\) The following iCn3D shows the interaction of the purified sodium channel with n-dodecyl-β-D-maltopyranoside (BDDM).  Explain the differences between the iCn3D models showing the protein in a bilayer and interacting with the BDDM. Answer . Exercise \(4\) The elution of the complex on a size exclusion column (Panel A) and the analyses for the eluted fractions by SDS-PAGE (Panel B) are shown in the figure below.  The α-scorpion toxin LqhIII: rat cardiac sodium channel NaV1.5 complex elutes in the area of the first peak shown in blue a.  How do molecules separate on size exclusion chromatography? b. Compare the molecular weights of the first peak to the second complex peak in Panel A. c.  Which band(s) in Panel B likely represent the rat cardiac sodium channel NaV1.5 based on the intensity of the stained band?  The toxin LqhIII is the lowest band.   ( bands at 17.5K and 12.5 are  FGF12b and calmodulin, respectively, which were added to stabilize the channel) d.  Why do the channel and toxin elute together in the size exclusion column shown in Panel A, but are separate bands in PAGE gel in Panel B? e.  How could you get the complex to separate as two peaks, the free NaV1.5 channel, and the free α-scorpion toxin LqhIII? f.  To get information on the receptor, go to Uniprot and paste in rat cardiac sodium channel NaV1.5 into the search box. Go to Sequence and Isoform in the left panel and find the actual MW.  Knowing this, what are the major bands at about 170K and 60K? Figure: Purification of the recombinant NaV1.5C/LqhIII complex. a. Representative size-exclusion chromatography profile of purified rNav1.5C/LqhIII. Peak fractions collected for cryo-EM grid preparation are shown in blue. b. SDS-PAGE of the size exclusion peak fractions stained by Coomassie blue. Answer . CryoEM was used to determine the structure of the sodium channel:toxin complex.  Here is a short YouTube video that describes the technique. Also review the appropriate part of Chapter 3.3: Analyses and structural predictions of protein structure. Exercise \(5\) Describe the temperature conditions for protein samples in cryoEM.  What is the reported resolution of cryo EM structure?  How does this compare to X-ray structures? What are some advantages of using cryoEM over X-ray crystallography and NMR to determine the structure of proteins? Answer . Molecular dynamics was also used to probe the conformational changes in the structure of the complex on the picosecond (10-12 s) to nanosecond (10-9 s) time scale.  For a review of molecular dynamics, see Chapter 3.3: Analyses and structural predictions of protein structure.  It can be used to probe dynamic changes in protein structure which cryoEM can't. Exercise \(6\) Answers these multiple choice questions (created by AIPDF through ChatGPT4 -paid version using this prompt:  Write 5 question for a biochemistry major about the use of molecular dynamics and the finding in the paper) 1. What was one of the primary uses of molecular dynamics in this research? -  A) Predicting the behavior of NaV channels without toxins. -  B) Analyzing hydration and Na+ permeation through the rNaV1.5C/LqhIII complex. - C) Studying the interaction between different toxins. - D) Predicting the behavior of potassium channels. 2. In the molecular dynamics simulation analysis, what was aligned to the initial position for each snapshot? - A) The α-toxin LqhIII. - B) The voltage-sensing domain IV. - C) The Cα atoms from pore transmembrane helices. - D) The fast inactivation gate. 3.  Approximately how long were the unrestrained "production" simulations generated? - A) 10.35 ns. - B) 5000 steps. - C) 300 ns. - D) 2 fs. 4. Based on the molecular dynamics analyses, what was observed about the activation gate structure of the rNaV1.5C/LqhIII complex? - A) It was fully open for Na+ conductance. - B) It was functionally closed for Na+ conductance. - C) It was in a metastable state. - D) It showed no significant change from the rNaV1.5C structure. Answer . The authors used two types of electrophysiological techniques, patch clamp, and voltage clamp.  Here is some brief background. In a whole-cell patch clamp experiment, a pipet is placed on a cell, and suction is applied until a tight seal, indicated by a sharp rise in electrical resistance (gigaohm level) is made.  This is illustrated in the figure below. Patch Clamp Resistance.  Formation of gigaseal.  Holst.  https://en.wikipedia.org/wiki/Automa..._Animation.gif.  CC BY-SA 3.0 The cell can then be connected to a patch clamp chip in such a way that transmembrane potential or current can be measured on single-channel ion flow. This is illustrated in the figure below. Holst. Patch Clamp Chip.  Batch clamp chip showing a gigaseal, whole-cell recording configuration, and the ion channel and whole cell current.  https://en.wikipedia.org/wiki/Automated_patch_clamp#/media/File:Patch_Clamp_Chip.svg.  CC BY-SA 3.0 In patch-clamp fluorometry, part of the cell membrane is sucked into the tip with the seal intact.  Fluorescent ligands can be applied to one side of the membrane that contains an ion channel and current measurements were made as illustrated in the figure below. Alternatively, as in this paper, side chains in the S4 voltage sensor were labeled with a fluorophore, and changes in fluorescence were observed with changes in membrane potential. Patch-Clamp Fluorometry.  https://www.uniklinikum-jena.de/phys...n/Methods.html Exercise \(7\) Answers these general multiple-choice questions (created by AIPDF through ChatGPT4 -paid version using this prompt: Write five multiple-choice questions about the use of patch clamp techniques to measure sodium currents in cells) 1. What is the primary purpose of the patch-clamp technique in cellular electrophysiology? - A) To visualize cell structures. - B) To measure the concentration of sodium ions inside cells. - C) To record ion currents across cell membranes. - D) To stimulate cellular growth. 3. In a typical neuron at resting potential (-70 mV) and in this study (epithelial cells transformed with the rat channel, what is the direction of the sodium current when sodium channels open? - A) Inward, into the cell. - B) Outward, out of the cell. - C) There is no movement of sodium. - D) Both inward and outward simultaneously. 4. Which of the following factors can influence the magnitude and direction of sodium currents measured using patch-clamp techniques? - A) The concentration of potassium ions outside the cell. - B) The voltage across the cell membrane. - C) The pH of the cell cytoplasm. - D) The size of the cell. 5. Why might a researcher use drugs or toxins during a patch-clamp experiment measuring sodium currents? - A) To increase the size of the cell. - B) To modulate or block sodium channels and observe the effects. - C) To change the color of the cell. - D) To stimulate cell division. Answer . Exercise \(8\) Answer these general multiple-choice questions about patch-clamp fluorometry. (created by AIPDF through ChatGPT4 -paid version using this prompt: write 5 multiple choice questions of patch clamp fluorometry in which key amino acids in a membrane protein are labeled with a fluorophore) 1. What is the primary advantage of combining patch-clamp with fluorometry in studying membrane proteins? - A) It allows simultaneous measurement of electrical activity and conformational changes. - B) It increases the fluorescence of all amino acids. - C) It enhances the electrical activity of the protein. - D) It allows visualization of the entire cell in detail. 2. Why are specific amino acids in a membrane protein labeled with a fluorophore in patch-clamp fluorometry? - A) To increase the size of the protein. - B) To change the electrical properties of the protein. - C) To detect specific conformational changes in the protein during activity. - D) To make the protein more soluble in water. 3. Which property of the fluorophore is crucial for patch-clamp fluorometry? - A) Its electrical charge. - B) Its sensitivity to changes in the local environment or protein conformation. - C) Its ability to increase protein activity. - D) Its color in visible light. 4. In which scenario would patch-clamp fluorometry be especially useful? - A) When studying the overall shape of a cell. - B) When investigating the relationship between ion channel gating and conformational changes. - C) When trying to increase the fluorescence of a solution. - D) When observing the movement of proteins inside the cell. 5. What is a critical consideration when choosing a fluorophore for labeling amino acids in patch-clamp fluorometry? - A) The taste of the fluorophore. - B) The electrical conductivity of the fluorophore. - C) The photostability and brightness of the fluorophore. - D) The size of the fluorophore molecule. Answer . Nonstructural Lab Studies of LqhIII Toxin Effects on Rat Sodium Channel NaV1.5 (rNaV1.5C) HEK293S GnTI (epithelial-like) cells were transformed with the rat cardiac sodium channel NaV1.5 (rNaV1.5C).  The cells were then studied in the absence and presence of the toxin at varying times after toxin addition and at various concentrations of the toxin. The opening and closing of the channel were determined by measuring changes in the Na+ currents into the cell on channel opening. Exercise \(9\) The resting potential of a cell is around -70 mV (more negative inside). When the transmembrane potential is depolarized by raising the transmembrane potential to around -55 mV or even more positive, the Na+ channels are activated, and an inward Na+ current (black line in a modified form of Figure 1a from the paper below) which goes downward by convention) through the channel occurs.  This is followed by a quick inactivation of the channel and the return to the baseline flow of ions.  In the experiment below, the potential was raised from -100 mV (channel closed) to 0 mV (channel open).  What is happening to the NaV1.5 Na+ channel during these 10 ms?  What is special about the current at 6 ms (indicated by the dashed vertical line) Answer Initially, the change in voltage opens the rNaV1.5C and allows an inward flow of Naions as evidenced by the vertical drop.  As described in the introduction, conformational changes in the NaV1.5C (closing of the inactivation loop), follow which closes the channel and stops the current, so the current returns to baseline within about 6 ms.  The channel is in the inactive state by around 6 ms. Exercise \(10\) Figure 1a from the paper (modified) below shows a series of lines of different colored (black to red) representing Na+ currents obtained at 0 (black line) and increasing concentrations (gray through red) of the LqhIII scorpion toxin.  Let's assume that the downward Ipeak =1.  The values of I 6ms/Ipeak, calculated from the approximate values shown on the graph,  are also shown on the vertical axis  Does the toxin alter the immediate response of the cells after the channel was activated?  What effect does increasing [toxin] have on the response of the cell?  Offer a structural explanation of how the toxin affects the cell by suggesting changes in the toxin-bound structure. Answer The toxin at any dose does not substantially affect the initial opening of the channel since the size of the current does not change at first.  Then the toxin inhibits the rapid inactivation of the channel within the first 6 ms, leading to a prolonged inward Na+ current.  The inhibition of the inactivation is dose-dependent on the concentration of the toxin. The toxin does not completely block the quick inactivation of the channel as the lines don't return to the black baseline in the time interval measured.  This suggests that the toxin binds to the channel and either partially occludes it or prevents the normal conformational change in the NaV1.5C that causes a quick deactivation of the channel. Since Na+ ions still flow through the channel but at a lower rate than the open state, the toxin-bound channel likely represents a 4th, or partially-opened state of the channel. Exercise \(11\) Figure 1a (left) from the paper below shows the dependency of the inhibition of the quick inactivation of the channel on the log of the LqhIII concentration.  When the transmembrane potential is set to 0 mV (as in this experiment), the channel should open and the current would be maximal.  The data points in the graph are close to the ones estimated in the graph from the previous question. a.  In the absence of the toxin, what should the current I be at 6 ms compared to the maximal Na+ current? That is, what would be the value of I6ms/Ipeak? b.  Is the channel completely inactivated in the presence of the toxin? c.  At 6 ms,  what concentration of toxin (nM) causes 50% inhibition of the maximum effect of the inhibitor on the normal rapid inactivation of the channel? Answer . Exercise \(12\) In the next experiment, cells were kept at -120 mV at one fixed concentration (100 nM) of toxin.  The toxin was left to incubate with the cells for various times up to 20 min.  After the incubation time, the transmembrane potential was changed to 0 mV to activate the channel, and inward Na+ currents were measured.  The results are shown in the top inset graph in Figure 1b from the paper.  (Note: It is unclear from the paper if the control was determined at 0 min with 100 nM toxin or no toxin.) a.  Did the length of time cells were pre-incubated with the 100 nM toxin affect Na+ currents after depolarization of the cells?  How did the effects on the cells depend on the preincubation time? b.  Describe and explain these results Answer . Here is another interesting feature of toxin binding.  The toxin binds to a site on the resting state of the NaV1.5C with high affinity.  When the cell becomes depolarized (made more + inside the cell), the affinity for the toxin decreases so it starts to dissociate. The affinity of the toxin for NaV1.5C decreases with increasing + transmembrane potential.  At very high positive potentials (+100 mV) it appears not to bind. Exercise \(13\) What might account for the decreasing affinity of the bound toxin for the  NaV1.5C with an increasing transmembrane potential? Answer . Exercise \(14\) Time course experiments were conducted on the complex at 100 nM of LqhIII scorpion toxin.  A three-pulse protocol can be applied to alter membrane potentials: • 1st: a pulse from −120 mV to +100 mV for the indicated times then • 2nd: a 50-ms hyperpolarizing pulse (make membrane potential very negative, perhaps around -100 mV) • 3rd: a pulse of 50 ms to 0 mV Note that steps 2 and 3 both occur with 0.1 s.  What is the purpose of each pulse? Answer . Exercise \(15\) Figure 1c from the paper below shows results for a set of 3-pulse designed to allow recovery of the fast inactivation which was blocked by the previous toxin binding.  Note that the transmembrane potential for the first pulse was +100 mV a.  Describe what happens to the channel and complex. b.  Explain the results c.  Summarize thermodynamic and kinetic features that make the toxin so effective. Answer . Structural Studies of LqhIII Toxin Effects on Rat Sodium Channel NaV1.5 (rNaV1.5C) - CryoEM Before we discuss in detail the structure of the sodium channel and its complex with the toxin, let's look at an important attribute of molecules that helps determine their function, the actual size of the species involved. Exercise \(16\) The Na+ and the K+ voltage-gated ion channels must have an open pore when the channel protein is active (Naions move across the channel).  Extracellular Na+ and the K+ ions don't exist as "naked" ions but they are hydrated by water in an aqueous extracellular environment. The figure below shows the relative sizes of these Group I cations and their hydrated forms, in comparison to the diameter of the open NaV1.5 pore in the channel.  Answer the following questions.  The red sphere (c) represents the calculated value of the diameter of water assuming its volume when it is bound to a protein is 25 Å3. 1.  Which represents the "naked" (nonhydrated) size of the K+ ion? 2.  Which represents the hydrated Na+ ion? 3.  Based on the pore size alone, which of these species could diffuse through the pore? Answer . Exercise \(17\) The approximate relative sizes of the hydrated Na+ ion, pore opening, toxin, and the NaV1.5 protein are shown in the figure below along with the relative width of the bilayer (BL).  A cardiac epithelium cell is shown as a rectangle to the right.  A red dot   (not visible in the large figure) in the membrane surrounded by the red-dotted circle represents a single NaV1.5 channel. 1.  Which likely represents the pore? 2.  Which represents NaV1.5 channel protein? 3.  Which represents the toxin? Answer . Exercise \(18\) If you hadn't read the paper, where would you consider the most likely location for a toxin to bind to affect the function of Nav1.5? Circle the most likely toxin binding site in the schematic below.  Based on the paper, where did it bind?  Redraw the toxin in the correct location based on the paper. Answer . Exercise \(19\) How is this toxin (LqIII) different in terms of its binding location from other toxins that interfere with the functioning of Nav1.5?  What is the effect of the toxin on the channel and the symptoms of this venom? Answer . Exercise \(20\) What amino acids should be present in the S4 segment of Nav1.4 and why? Answer . Exercise \(21\) The figure below shows an interactive iCn3D model of the rat sodium channel NaV1.5 bound to the LqhIII toxin  (7k18). a.  To which domain does the LqhIII toxin bind? b.  Is the binding site close to the pore-forming segments of the domain or the voltage-sensitive segments? c.  Does the layer of red spheres represent the outer (extracellular) or inner (intracellular) leaflet of the membrane? d.  Offer reasons that parts of the protein are missing from the structure. Answer . Exercise \(22\) The IFM motif has been shown to be conserved across all voltage-gated sodium channels. a.  What role does it play in these channels and in Nav1.5 and why? b.  The cause of Paroxysomal Extreme Pain Disorder (PEPD), an extremely rare disease with only 15 known affected families, appears to be mutations in the IFM motif which leads to increased sensations of pain.  What is a likely effect of the mutations in the IFM motif? https://en.wikipedia.org/wiki/Paroxy..._pain_disorder Answer . Exercise \(23\) The figure below shows a different interactive iCn3D model of the rat sodium channel NaV1.5 bound to the LqhIII toxin (7k18) without a membrane representation for clarity.  It shows the selectivity filter DEKA (spacefill, CPK colors), the inactivation gate IFM and the IFM "internal receptor" F1651, L1660, and N1662 (spacefill, CPK colors), and a ring of hydrophobic residues V413, L941, I1471, and I1773 (spacefill, black) that in the closed state completely seal off the cytoplasmic opening in the pore. Rat sodium channel NaV1.5 bound to the LqhIII toxin without a membrane representation (7k18). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...pPzakHu6Ew6WH9 After viewing the structure from all angles, do you think that the toxin:NaV1.5 complex looks closed, open, or inactivated form?  Explain Answer . Structural Studies of LqhIII Toxin Effects on Rat Sodium Channel NaV1.5 (rNaV1.5C) - Molecular Dynamic Simulations Exercise \(24\) Now let's look at some data to see if the pore is really open, partially open, or closed in the toxin:channel complex. One clue is if the structure shows water in the pore as the Na+ ions must be hydrated to pass through the pore (the opposite case is seen with K+ channels when K+ pass through stripped of water).  Molecular dynamic (MD) simulations were done on the NaV1.5 with and without the toxin to simulate the environment in the channel opening.  The results of the MD simulations are shown below in Figures 6 a and c from the paper. Molecular dynamics analysis of hydration and Na+ permeation through the rNaV1.5C/LqhIII complex. Panel a shows a side view of rNaV1.5C (orange ribbons; domains II and IV) from MD simulations highlighting Na+ ions (blue spheres), the water-occupied volume within a cylinder of radius 8.5 Å (red surface), and the protein-occupied volume within a cylinder of radius 12 Å (colorless surface). The cavity within the pore is outlined with a black rectangle. The region of the intracellular activation gate is shown as a purple band. Panel c shows molecular representations of the gate containing Nwater = 3 (left) or 15 (right) water molecules Based on these studies, do you believe the pore is closed, open, or partially open? Answer Detailed Structural Analyses of LqhIII Toxin Effects on Rat Sodium Channel NaV1.5 (rNaV1.5C) Exercise \(25\) From the iCn3D model, write the sequence of the S4 segment that contains the Arg side chains and describe the properties of the amino acids in the sequence.   Do this by scrolling along the sequence window in iCn3D (shown below) until you find the labeled Arg shown in the model. Answer . Exercise \(26\) Is the helix amphiphilic?  That is, are the Arg side chains all on one face of the helix and the nonpolar on the other?  To find out, copy and paste the sequence of S4 (above) in this helical wheel predictor and run the program. Answer . Exercise \(27\) Make a simplified view of the iCn3D model by hiding Domains 1-III to more readily see the contributions of S5 and S6 of Domain IV to the pore. • open iCn3D and load 7K18. • With your mouse or trackpad, choose Sequence and Annotation in the top menu bar • Choose the Details tab • Ctrl-Click the two sequences highlighted in yellow below for Domain IV and the toxin. • Choose View from the top menu bar and then View Selection • Choose Style, Background, Transparent . Noncovalent Interactions of LqhIII and Domain IV/VS Now let's look at the actual interaction of the toxin with the Domain IV/VS of the channel.  A closeup showing the interaction site is shown in Figures 3 b and c from the paper below.  Panel C next to it shows the NMR-solution structures of the toxin in the absence of the channel.   Each structure determined is represented by a single color line color codes red at the C-terminus to blue at the N-terminus. Panel b: CryoEM structure of the  rNaV1.5C Domain IV/VS and LqhIII complex; Panel c:  NMR structure of free LqhIII Exercise \(28\) Using this iCn3D model, describe the secondary structure of the bound toxin.  How many pairs of cysteine residues are in the LqhIII toxin. Identify which cysteines are involved in the disulfide bonds.  What effect do the disulfide bonds have on the beta sheet structure? Answer . Exercise \(1\) What sections of the toxin in panel B make the closest interactions with the Domain IV/VS of the channel?   Describe their conformation flexibility in the free toxin. Answer . The figure below shows an interactive iCn3D model of a surface rending of Domain IV of the rat sodium channel NaV1.5 bound to the LqhIII toxin  (7k18). Surface rending of Domain IV of the rat sodium channel NaV1.5 bound to the LqhIII toxin  (7k18). (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...x5QRPEctT5zsW7. The molecular surface and underlying secondary structure of the LqhIII toxin are shown in magenta, with key residues H15, H43, and K64 shown as CPK-colored sticks and labeled.  Domain IV/VS of the channel is shown in cyan, with key amino acid side chains T1608, D1612, and Q1615 shown as colored and labeled sticks. Exercise \(29\) Comment on the shape and possible side chain interactions that contribute to high-affinity binding of the inhibitor to Domain IV/VS. Answer . Figure 3d from the paper below shows the detailed interactions between LqhIII and DIV-VS. Key residues shown in sticks were labeled. Interaction surfaces of the DIV-VS (blue) and the LqhIII (purple). Key residues for the interaction are shown in yellow shading and embedded stick The figure below shows an interactive iCn3D model of the rat sodium channel NaV1.5 Domain IV bound to the LqhIII toxin  (7k18). The toxin is shown in magenta.  The segments are colored as follows: S1 is red, S2 is orange, S3 is yellow, S4 is cyan, S5 is brown and S6 is violet.  Key amino acid pairs involved in the binding of the toxin to Domain IV are shown in sticks and labeled. Exercise \(30\) In summary, name and locate the amino acid residues that serve the following roles in the LhqIII toxin: DIV-VS interactions. 1. Which amino acids in the toxin interact with D1612 (the paper describes the interaction as pincers surrounding D1612). 2. The conserved negatively charged residue in the Nav1.5 channel 3. What position is Thr in and what is thought to be its role in the mechanism? Answer Comparison of Activated DomIV-Voltage sensor (VS) with Toxin-bound Partially activated DomIV-VS Now we'll try to understand Figure 4, Conformational Change of DIV-VS,  from the paper and pay special attention to the section of the text, “An intermediate-activated state of DIV-VS trapped by LqhIII” - Let’s dissect Figure 4 A and B. Figure 4a/4b from the paper below shows the conformational change of Domain IV-Voltage sensor (DIV-VS) comparing the activated and partially activated state with the bound toxin. Panel a shows the activated domain IV with key Arg side chains in S4.  Panel B shows the partially activated domain IV with the same key Arg side chains in S4.  The bound LqhIII is shown as a purple chain.  In panels a and b, the: 1. activated Nav1.5 DIV-VS (the voltage sensing domain) is in grey (fig 4a) 2. the intermediate-activated Nav1.5DIV-VS is in blue (Fig 4b) 3. side chains of gating charges of Arg are shown in grey and blue sticks in 4a and in shades of blue sticks in 4b. Side chains in the ENC are shown in red, in the HCS in yellow, and in the INC  in red; 4. the shift of each gating charge was indicated by black dashed lines between the structures in panels a and b. The shift from the cytoplasmic to extracellular parts of the channel is shown in the region between the two panels. The black Rs in the activated DIV-VS(panel A) are further up in the diagram (towards the extracellular region) and further down in the partially activated DIV-VS bound to the toxin. Exercise \(31\) Locate the 6 arginines? (R1-R6) with the blue indicating the N atoms in the positively charged Arg side chain of S4 in Domain IV in one of the iCn3D models above.   What do the following abbreviations mean? ENC, HCS, and INC. Answer . Exercise \(29\) From Figure 4a to 4b, explain from an electrostatic viewpoint how the movement of the Args towards the extracellular region would promote the movement of Na ions inward.  Explain how the movement of Na+ ions would be diminished in the presence of the toxin. Answer . Comparison of Active and Intermediate-Activated, and Intermediate/Resting state Now consider Figures 4 C and D from the paper below: 4c: Superposition of NaV1.5 DIV-VS between the fully activated state and toxin-bound intermediate-activated state. Red arrows indicate the conformational changes. 4d: Superposition of the intermediate-activated NaV1.5 DIV-VS and resting-state NaVAb-VS Exercise \(32\) a.  Locate the region in Figure c above that shifts the most from the fully activated to intermediate-activated state of the DIV-VS. b.  Describe the difference shown in Figure d between the intermediate-activated DIV-VS structure (blue) upon the resting state NaVAb-VS structure (orange) c.  What do these differences imply about the conformational states of the apo and toxin-bound channel? Answer . Summary Exercise \(33\) Why might the mode of action be specific for cardiac muscle cells as compared to other toxins that act on sodium channels in skeletal and nerve cells? Answer . After this guided research literature module, you can hopefully better understand the findings in the paper which are summarized in this abstract: "Voltage-gated sodium (NaV) channels initiate action potentials in excitable cells, and their function is altered by potent gating-modifier toxins. The α-toxin LqhIII from the deathstalker scorpion inhibits fast inactivation of cardiac NaV1.5 channels with IC50 = 11.4 nM. Here we reveal the structure of LqhIII bound to NaV1.5 at 3.3 Å resolution by cryo-EM. LqhIII anchors on top of voltage-sensing domain IV, wedged between the S1-S2 and S3-S4 linkers, which traps the gating charges of the S4 segment in a unique intermediate-activated state stabilized by four ion-pairs. This conformational change is propagated inward to weaken binding of the fast inactivation gate and favor opening the activation gate. However, these changes do not permit Na+ permeation, revealing why LqhIII slows inactivation of NaV channels but does not open them. Our results provide important insights into the structural basis for gating-modifier toxin binding, voltage-sensor trapping, and fast inactivation of NaV channels." Extensions 1.  Interesting sidebar:  https://www.sciencedirect.com/science/article/pii/S0021925819308300?via%3Dihub Chlorotoxin (Cltx) is a 36-amino acid peptide that was originally isolated from Leiurus quinquestriatus venom (14) and has been shown to inhibit small conductance Cl− channels in colonic epithelial cells (14, 15). Cltx also inhibits Cl− fluxes across glioma membranes (13, 16). Immunohistochemical studies show that Cltx specifically and selectively binds to glioma cells (17) and radiolabeled Cltx targets tumor cells in mice bearing xenografted glioma tumors. Glioma cell migration and invasion into fetal brain aggregates is significantly reduced by Cltx (13). A recent survey of over 200 tissue biopsies from patients with various malignancies suggests that Cltx binds to the surface of gliomas and other embryologically related tumors of neuroectodermal origin (18) but not to normal brain. 2.  Deathstalker scorpion venom also contains chlorotoxin - This is a very interesting story. Note and remember LqhIII is an alpha toxin 4.  How much is deathstalker venom worth? 5.  Looks like a great review article: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC7277529/ 6.  Scorpion Venom: Detriments and Benefits 7.  Lookfor possible therapeutic potential here https://www.venomdoc.com/ 8.  Very cool venom graphic - https://www.venomdoc.com/new-page-2 9.  The Toxicogenomic Multiverse: Convergent Recruitment of Proteins Into Animal Venoms:  https://static1.squarespace.com/static/55a239e2e4b0b3a7ae106f25/t/59814ceae6f2e10bc7ada5f1/1501646072821/2009_Fry_Toxicogenomic_multiverse.pdf 10.  The deathstalker scorpion venom alone has been found to have several different kinds of toxins including chlorotoxin (inhibit chloride channels), charybdotoxin (inhibit potassium channels), and agitoxins (affect sodium channels)..  https://www.sciencedirect.com/topics/biochemistry-genetics-and-molecular-biology/leiurus-quinquestriatus. Chlorotoxin was found to selectively bind to glioma cells and serve as a marker for glioblastoma.  https://www.acs.org/molecule-of-the-week/archive/c/chlorotoxin.html#:~:text=Strichartz%20at%20Harvard%20Medical%20School,diagnosing%20and%20treating%20some%20cancers. This feature of the scorpion venom was developed by J.M. Olson at Fred Hutchinson Cancer Center (Seattle) as a product called Tumor Paint https://www.fredhutch.org/en/news/center-news/2014/09/tumor-paint-US-trial.html

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/Structure%2F%2FFunction_-_Protein_Problems/LGA%3A_Voltage-Gated_Sodium_Channel_-_Students_082423.txt

Carbonic Anhydrase Engineered Stability We have already encountered this enzyme before (Chapter 6.1). It catalyzes the hydration of CO2 (g) as shown below. CO2 (g) + H2O ↔ H2CO3 (aq) ↔ HCO3- (aq) + H+ (aq) It is among the fastest of all enzymes, with a kcat of 106 s-1 and a kcat/Km of 8.3 x 107 M-1s-1 (reference). It is diffusion controlled in that the rate of diffusion of reactants and products, not the chemical steps, determine the reaction rate. It can convert 106 molecules of CO2(g) to HCO3- each second. No wonder scientists and engineers are studying it to capture CO2. It's a big challenge though to capture CO2 released on combustion of coal or natural gas in a power plant. Here are two problems that must be overcome: • The enzyme must be thermostable at elevated temperatures to capture the CO2 found in high-temperature power plant emissions • The enzyme is reversible so it will be inhibited by the product HCO3- • The enzyme must be stable to somewhat alkaline conditions (pH of 0.1M NaHCO3 = 8.3) For carbon capture from fossil fuel emissions, CA is immobilized by surface adsorption, covalent attachment, encapsulation, and entanglement. Immobilized enzymes are typically more thermostable and can be used in flow-through as opposed to solution phase capture. The immobilized enzyme matrix must withstand high temperatures (up to 100°C, and alkaline solvents used to strip the matrix for reuse. The enzyme is found throughout life and typically has an active site Zn2+. There are 8 families, α, β, γ, δ, ζ η, θ, and ι, with the α family being the most abundant. The α forms are generally active as dimers, but can act as monomers and tetramers.. There are 15 isoforms of the α form in humans and have a prime role in pH regulation. They are found in bacteria, fungi, plants, and algae. β-CAs are found in some types of bacteria, archaea, fungi, some higher plants, and invertebrates. CA in chloroplasts (and mitochondria (algae) are involved in carbon fixation. We will focus our attention on engineering carbonic anhydrase to make them more thermostable, alkali insensitive, and less susceptible to product inhibition by bicarbonate. Natural enzymes can be isolated and selected for thermal and alkali stability. In addition, new versions selected for these properties can be engineered using directed evolution or site-directed mutagenesis. You wish to increase the thermal stability of a protein using mutagenesis. Essentially you wish to perturb the equilibrium between the folded (native) protein and the unfolded (denatured) protein so as to preferentially stabilized the native state. Question $1$ Using mutagenesis, what residues might you change in a native protein to make it more stable at higher temperatures? Answer A characteristic of the native state of the protein is its conformational stability compared to the conformational flexibility of the many possible denatured states. In addition, the protein must undergo conformational changes as it unfolds. Hence anything that restricts conformational flexibility might preferentially stabilize the native state. These would include changing single or pairs of side chains to allow the formation of more salt bridges and intrachain disulfide bonds, as well as hydrogen bonds. Loops with greater flexibility, as determined by B-factors in the crystal structure files, or by molecular dynamic simulations, could be changed to contain a disulfide, which would clearly stabilize a flexible loop. Question $2$ What measurements would you make to quantitate the change in thermal stability? Answer Measures a signal that changes with increasing temperature. The signal can be enzyme activity, or more easily a spectroscopic signal such as absorbance at 280 nm or fluorescence as a function of temperature. Alternatively, the stability at room temperature could be measured using urea as a perturbant. These are discussed in Chapter 4.12. The actual amino acid composition and more strangely specific dipeptide sequences within a sequence are associated with thermal stability of hyperthermophilic proteins. For example, proteins from two different types of archaea with different optimal growth temperatures show that the one with the higher growth temperature have significantly higher levels of VK, KI, YK, IK, KV, KY, and EV and decreased levels of DA, AD, TD, DD, DT, HD, DH, DR, and DG. Similar experiments have been done in bacterial cells. Using machine learning, the dipeptide sequences KH, KR, TF, PM, F∗∗N, V∗∗Y, MW, and WQ were important in themostability where the * denotes a gap in the residues. Carbonic anhydrase from Neisseria gonorrhea (ngCA) Data from: Jo, B., Park, T., Park, H. et al. Engineering de novo disulfide bond in bacterial α-type carbonic anhydrase for thermostable carbon sequestration. Sci Rep 6, 29322 (2016). https://doi.org/10.1038/srep29322. Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/ Now that we understand the general chemistry, structure, and reaction mechanism of carbon anhydrase (at least the alpha human CAII form), let's explore efforts to engineer more thermostable variants. One example is the carbonic anhydrase from N. gonorrheas. This CA has been used as a target for mutagenesis to increase thermal stability of the enzyme, through the introduction of new disulfide bonds. Even though only about 35% of the amino acids are identical, the overall structures are similar. This is illustrated in Figure $10$ below. Figure $10$: Alignment of the carbonic anhydrase from Neisseria gonorrhea (NG-CA) magenta,1KOQ) and human CA II (cyan, 2VVB) The active site is mostly conserved compared to human CA II. The Zn2+ bound water has a pKa of around 6.5, compared to the value of 7.0 in human CA II. The hydrophobic patch (pocket) is similar, with Phe 93, Leu 153 and Tyr 72 in the NG-CA replacing Phe 95, Phe 176, Phe 70 in human CA II, respectively. The histidine ligands to Zn2+ are His92 (94), His94 (96), and His111 (119), where the numbers in parentheses represent Hu CA II. The proton removed from Zn2+ bound water is transferred to His 66 (64 in human CA II) and then to His 64. The single disulfide bond between 181 and C28 is shown in Figure $1$ below Figure $11$: Single disulfide bond between 181 and C28 in wild type Carbonic anhydrase from Neisseria gonorrhea Question $18$ This question addresses the Biomolecular Visualization Framework theme(s) Atomic Geometry (AG) Identify the correct torsion angles in Figure $\PageIndex{x}$ above. Verbal definitions of torsional angles in a peptide chain are listed below. The successive atoms after the Cα leading away from the backbone atoms are Xβ-Xγ-Xδ-Xε (in that order). C is the backbone carbonyl C and N is the backbone nitrogen atom. • phi (φ) is the angle of right-handed rotation around N-Cα bond. φ = 0 if the Cα-C bond is cis (eclipsed) to the C-N bond. Values range from -180 to 180 degrees. • psi (ψ) is the angle of right-handed rotation around Cα -C bond. ψ = 0 if the C-N bond is cis (eclipsed) to the N-Cα bond. Values range from -180 to 180 degrees. • chi11) is the rotation around N-Cα-Xβ-Xγ • chi22) is the rotation around Cα-Xβ-Xγ-Xδ • chi33) is the rotation around Xβ-Xγ-Xδ-Xε Answer phi (Φ) = b, psi (Ψ) = a, chi11) = c, chi22) = d, and chi33) = e Question $19$ This question addresses the Biomolecular Visualization Framework theme(s) Atomic Geometry (AG), Topology and Connectivity (TC) Figure $12$ below shows an interactive iCn3D model of the atoms within 4A of the disulfide bond in Carbonic anhydrase from Neisseria gonorrhea (1KOQ). Rotate the model to determine the approximate chi33) dihedral angle. Hint: site down the S-S bond. Figure $12$: Atoms within 4A of the disulfide bond in Carbonic anhydrase from Neisseria gonorrhea (1KOQ). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...nutjP6EL2PubRA Answer The visually estimated chi33) angle for rotation around the S-S bond is 90o. Here is the actual angle (image made with Pymol) We mentioned previously that variants with higher thermal stability are likely to be more rigid and less flexible. Flexibility can be determined through analysis of molecular dynamic simulations and also by examining of the B factor values in PDB file. This number is a measure of the displacement of an atom from a mean The numbers in the last column in the file are called the temperature factors or B-factor. The B-factor describes the mean-square displacement, a measure of the displacement of an atom from an average value. If the atoms are more flexible, the electron density determined in x-ray structures is lower than if the atoms are more fixed, which gives high electron density. To make stabilizing disulfide bonds, investigators found site chains close enough that when mutated to cysteines could potentially form disulfide bonds. In addition, they search for such residues in surface loops (without alpha and beta structure) which are inherently more flexible. Introducing disulfide bonds into the loop would stabilize it and make it more rigid. Table $2$ below shows a description of the double cysteine CA variants in the study. Variant designation Position Wild-type residues Loop length Sum of B-factors T133C/D197C 133, 197 Thr/Asp 63 87.60 P56C/P156C 56, 156 Pro/Pro 99 80.82 N63C/P145C 63, 145 Asn/Pro 81 77.17 Table $2$: Description of the double cysteine CA variants in ngCA The locations of the side chair targeted for mutations to cysteine pairs are shown in Figure $13$ below. Figure $13$: 3D structure of ngCA and location of residue pairs for disulfide engineering The zinc (not shown)-coordinating histidine residues in the catalytic active site are shown in green. The proton shuttle histidine residue is shown in magenta. The native disulfide bond is colored yellow. An interactive iCn3D model of carbonic anhydrase from Neisseria gonorrhea (1KOQ) highlighting the 3 pairs of sidechains for mutations is shown in Figure $14$ below. Figure $14$: Carbonic anhydrase (Neisseria gonorrhea) with 3 paired side chains for engineered disulfide (1KOQ) (Copyright; author via source). Click the image for a popup or use this external link: https://structure.ncbi.nlm.nih.gov/i...KRP2tFJ1pV1FF6 The mutations were made and the wild-type proteins and three mutants were subjected to SDS-polyacrylamide gel electrophoresis (SDS-PAGE). The stained gels are shown in Figure $15$ below. Figure $15$: Expression and purification of disulfide CA variants. Panel (a) shows the expression of the protein in transformed cells 25 °C after IPTG induction and fractionated into soluble and insoluble fractions. SHuffle strain, an engineered E. coli strain that promotes cytoplasmic disulfide bond formation, was used. Panel (b) shows purification results. Each lane was loaded with 4 μg of each purified CA variant. The proteins were visualized with Coomassie blue staining after SDS-PAGE. The arrow indicates the position of the bands corresponding to ngCA variants. Lane: M, molecular weight marker; S, soluble fraction; IS, insoluble fraction. Question $20$ a. Interpret the results of the PAGE gels in Figure $15$ above b. How pure were the proteins based on the PAGE gel result in Panel (b). Can you infer from the gel that the proteins folded correctly? Answer a. It appears to show that a small fraction of the wild type and each mutant, especially the N63C/P145C pair, was found in the insoluble fraction. This might result from improper folding of the proteins in E. Coli, leading to hydrophobic side chain surface exposure and aggregation into insoluble "inclusion bodies". This appears to be just a minor issue. b. All the proteins appear very pure with some very small levels of contamination in the N63C/P145C. The PAGE results show the protein all have the same molecular weight but whether they folded to a native state with activity can not be determined. Nor can it be determined if the disulfide pairs in the mutants were made are if they were, were correctly paired. The investigators next determined if the expressed and purified wild-type and mutant proteins had the correct number of disulfide bonds. They did this by reacting the proteins in the absence and presence of dithiothreitol with DTNB or 5,5'-dithiobis(2-nitrobenzoic acid), also called Ellman's Reagent. Both structures are shown in Figure $16$ below. Figure $16$: Structures of DTT and DTNB DTT is a reducing agent that cleaves disulfide. Question $21$ Draw a mechanism showing the reaction of a disulfide with DTT. Answer Only surface and not buried free cysteines will be labeled unless the protein is unfolded to expose all the cysteines. DTNB reacts with free sulfhydryls to form the 2-nitro-5-thiobenzoic acid anion leaving group that absorbs at 412 nm. Question $22$ Draw a mechanism showing the reaction of free sulfhydryl (like Cys) with Ellman's reagent Answer Only surface and not buried free cysteines will be labeled unless the protein is unfolded to expose all the cysteines. The results of the reaction of the proteins with Ellmans's agent, in the presence and absence of DTT, are shown in Table $3$ below. CA variant Free thiol/protein (mol/mol) a Deduced no. S-S bonds −DTT +DTT Wild-type 0.06 ± 0.02 1.80 ± 0.14 ? T133C/D197C 0.08 ± 0.02 3.79 ± 0.22 ? P56C/P156C 0.06 ± 0.03 3.89 ± 0.06 ? N63C/P145C 0.08 ± 0.03 3.75 ± 0.05 ? Table $3$: Analysis of disulfides in CA using Ellman's reagent. aNumbers are represented in mean ± SD. Question $23$ How many S-S would you deduce from the table are present in the wild-type and mutant enzymes? Did the correct disulfide bonds form? Explain your answers Answer DTT reduces the disulfide in protein. For each disulfide, two free Cys side chains are made. The molar ratio of CysSH/CA for the wild-type is 1.8 in the presence of DTT. The value is very close to the expected value of 2. For the mutants, the ratio is about 3.8 in the presence of DTT, suggesting 4 free Cys consistent with 2 disulfide bonds. Table $4$ below shows the catalytic activities of the disulfide CA variants at 25 °C. CA variant CO2 hydration activity Relative esterase activity a kcat × 10−4 (s−1) K M (mM) kcat /KM × 10−6(M−1 s−1) Wild-type 1.00 1.44 14.2 1.01 T133C/D197C 1.49 1.97 16.7 1.18 P56C/P156C 1.03 1.44 16.9 0.85 N63C/P145C 0.55 0.27 17.3 0.16 Table $4$: catalytic activities of the disulfide CA variants at 25 °C aThe specific activity of the wild-type corresponds to 0.22 U/μmol-enzyme. Question $24$ Why did the investigators conduct this experiment? Interpret the results Answer All of the previous results suggest that the mutant proteins were made and had the correct number of double bonds, but the experiments could not tell if the bond pairs were correct. For example, did the T133C/D197C contain a native (C28-C181) and mutant (C133-C197) bond and not another combination? Activity is an excellent predictor of structure. All but one mutant retained nominal activity, as evidenced by a comparison of the rat constants. The N63C/P145C had a 6x lower kcat, but even then it is close to diffusion-controlled. Now comes the big question: were the investigators able to engineer thermal stability into the carbonic anhydrase? Experimental results to show the thermostability of the disulfide CA mutants are shown in Figure $17$ below. Figure $17$: Thermostability of the disulfide CA variants Panel (a) shows short-term kinetic stability. The enzyme solutions (40 μM) were incubated for 30 min at different temperatures, and the residual activities were measured by esterase activity assay. Activities of 100% correspond to untreated samples. Panel (b) shows long-term kinetic stability at 70 °C. The half-lives (t1/2) of the CA variants were estimated by fitting the experimental data to an exponential decay curve. Each value represents the mean of at least three independent experiments, and the error bars represent the standard deviations. Question $25$ Analyze the results in Panels (a) an (b). Which protein was most thermostable over the short (30 minute) and long (hours) incubating time at elevated temperatures? Answer At 80 °C, all the mutants showed increases thermostability to short term (30 minute) heating, but one, N63C/P145C was exceptionally thermostable. Longer time courses for heating at 70°C showed that the N63C/P145C was again far more stable over time. Its t1/2 was 31.4 h, compared to the values between 4-6 h for the others. Panel (c) shows heat-induced denaturation of disulfide CA variants. Temperature-dependent changes of the circular dichroism ellipticity were recorded at 220 nm on CD spectrometer. The denaturation curves were normalized to the fraction of unfolded protein. The horizontal dashed line indicates the point at which the fraction of unfolded protein is 0.5. The vertical dashed lines point to TM values. Panel (d) shows the overall RMSD of disulfide variants from molecular dynamic simulations performed at 400 K for 20 ns. Question $26$ Analyze the results in Panel (c). What do changes in the CD helicity show? Which protein was most thermostable based on TM values? Is the decrease in enzyme activity in panels (a) and (b) result from the denaturation of the protein? Answer CD measurements can give a measure of the retention of secondary structure (alpha helices and beta structure) on denaturation. The CD spectrum for different secondary structures is shown below (From Chapter 3.5). The curves were normalized to fit on a 0-1 scale on the y axis, which then gives a measure of percent denaturation. The temperature half-way up is the TM, or "melting temperature, at which an equilibrium mixture would contain half native and half denatured protein (true for a small protein with no intermediates). The TM values were for the wild-type, T133C/D197C, P56C/P156C, and N63C/P145C mutants 73.6 °C 74.7 °C, 77.4 °C, and 81.4 °C. These parallel the t1/2 values for enzyme activities, and support the idea that denaturation led to inactivation of the enzyme. Question $27$ Analyze the molecular dynamics simulation results in Panel (d) Answer The molecular dynamic simulations for all the proteins soon reach equilibrium values as indicated in the plateaus of average room mean square deviation of the protein backbone. The overall molecular root-mean-square deviation (RMSD) of N63C/P145C was the lowest, indicating that it was most rigid. This is in accord with the idea that increased flexibility destabilizes a protein and engineering a disulfide into makes it more rigid and hence more stable to temperature increases. The results are in accordance with the other experiments that show the N63C/P145C was the most thermostable. " In addition, T133C/D197C showed the highest values in both the overall and the residual RMSD (Fig 3D). This may explain and correlate with the increased activity of T133C/D197C (Ta) and the increased ΔS of unfolding" You may remember from both introductory chemistry and from Chapter 4.12, that you can calculate the thermodynamic parameters, ΔHo and ΔSo for N ↔D at room temperature from thermal denaturation curves using the van 't Hoff equation. In this case, Keq values can be calculated from thermal denaturation curves by monitoring change in CD signal at 220 nm, and applying this equation (also from Chapter 3.12). K_{e q}=\frac{[D]_{e q}}{[N]_{e q}}=\frac{f_D}{f_N}=\frac{f_D}{1-f_D} From this, we can calculate ΔG0. \Delta \mathrm{G}^0=-\mathrm{R} \operatorname{Tln} \mathrm{K}_{\mathrm{eq}}=-\mathrm{R} \operatorname{Tln}\left[\frac{\mathrm{f}_{\mathrm{D}}}{1-\mathrm{f}_{\mathrm{D}}}\right] Knowing Keq, ΔH0, DS0 can be calculated as shown below. A semi-log plot of lnKeq vs 1/T is a straight line with a slope of - ΔH0R and a y-intercept of + ΔS0/R, where R is the ideal gas constant. \begin{gathered} \Delta \mathrm{G}^{0}=\Delta \mathrm{H}^{0}-\mathrm{T} \Delta \mathrm{S}^{0}=-\mathrm{RTln} \mathrm{K}_{\mathrm{eq}} \ \ln \mathrm{K}_{\mathrm{eq}}=-\frac{\Delta \mathrm{H}^{0}-\mathrm{T} \Delta \mathrm{S}^{0}}{\mathrm{RT}} \ \ln \mathrm{K}_{\mathrm{eq}}=-\frac{\Delta \mathrm{H}^{0}}{\mathrm{RT}}+\frac{\Delta \mathrm{S}^{0}}{\mathrm{R}} \end{gathered} The equation below shows that the derivative of equation (8) with respect to 1/T (i.e. the slope of equation 8 plotted as lnKeq vs 1/T) is indeed -ΔH0/R. Equation (9) is the van 't Hoff equation, and the calculated value of the enthalpy change is termed the van 't Hoff enthalpy, ΔH0vHoff. \frac{d \ln \mathrm{K}_{\mathrm{eq}}}{d(1 / \mathrm{T})}=-\frac{\Delta \mathrm{H}^{0}}{\mathrm{R}}=-\frac{\Delta \mathrm{H}_{\mathrm{vHoff}}^{0}}{\mathrm{R}} Using this method, the thermodynamic parameters for unfolding of the protein were calculated. The results are shown in Table $4$ below. CA variant Melting temperature, TM (°C) Enthalpy change of unfolding, ΔH (kcal mol−1) Entropy change of unfolding, ΔS (kcal mol−1 K−1) Wild-type 73.6 48.8 0.141 T133C/D197C 74.7 52.8 0.153 P56C/P156C 77.4 35.1 0.091 N63C/P145C 81.4 30.0 0.085 Table $4$: Thermodynamic parameters for protein unfolding for WT and mutant CAs Question $28$ Which effects, enthalpy or entropy of unfolding, were associated with the increased thermal stability of the mutants compared to the wild-type protein. Remember were are considering the denaturation reaction, N↔ D. Answer For the reaction N ↔ D, the ΔH0 values were all positive, indicating the enthalpy changes favored the native state, not the denatured state. In contrast, the other two mutants were enthalpically destabilized compared to the wild-type as their ΔH0 were less positive so compared to the wild-type. The prime stabilizer of the native state was the lower entropy (hence a less negative and favored -TΔS0 for the denaturation reaction. This makes sense in these mutants are more rigid and would experience less loss of "conformational entropy). P56C/P156C and N63C/P145C exhibited lower ΔH (destabilizing) and ΔS (stabilizing), showing that the decreased entropic change of unfolding (i.e., the loss of conformational entropy of the unfolded state) by the disulfide bridge was the primary factor for the thermostabilization. These results are not surprising because design strategies aiming ‘entropic stabilization’ such as disulfide engineering do not always result in engineered proteins ideally with lower ΔS and unchanged ΔH. These results are in accord with the observation that N63C/P145C was the most thermostable variant and that T133C/D197C showed the highest values in both the overall and the residual RMSD (Fig. 3d). This may explain and correlate with the increased activity of T133C/D197C and the increased ΔS of unfolding. Finally, the enzymatic activity of the wild-type and mutants CAs (using a small ester substrate) were studied as a function of temperature. The relative activity of the wild-type and all 3 disulfide mutants are plotted as a function of temperature in the histogram graphs shown in Figure $18$ below. Figure $18$: Effect of temperature on the activity of disulfide CA variants. Esterase activities of disulfide variants were measured at each temperature and normalized to the activity of each enzyme at 25 °C. Each value represents the mean of three independent experiments, and the error bars represent the standard deviations. If you plotted the data as curves, you would get bell-shaped graphs. Question $29$ Explain why the histogram plots (and line plots if they were drawn) are bell-shaped. Are the results in accordance with the previous results. Answer Yes. Most chemical reaction show an increase in rate with increasing temperatures until competing reactions take precedence. For an enzyme-catalyzed reaction, that competing reaction is denaturation, which decreases the rate. Yes the graphs are in accord with the previous results. The N63C/P145C certainly stands out as the best mutant. The authors write that "considering the shifted optimal temperature and the thermoactivation as well as the enhanced thermostability, the disulfide engineered α-type CA with Cys63-Cys145 can be a promising biocatalyst for efficient CO2 sequestration performed under high temperature conditions."

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/Structure%2F%2FFunction_-_Protein_Problems/Literature-based_Guided_Assessments%3A__Protein_Stability_-_Carbonic.txt

File, Retrieve by ID MMDB • MMDB (Molecular Modeling Database) files from the NCBI • derived from PDB atomic coordinates but with … • Database information (quaternary struct, molecular interactions, SNPs, conserved domains,  clinical variants – i.e related structure info, not just xyz coord PDB • xyz coordinates RCSB MMTF ID (fast) • Great for very big structures that otherwise too slow in loading; Few modeling options. AlphaFold Structure • Computationally determined structures • Uses Uniprot or RCSB ID. OPM PDB ID • Get structures on membrane proteins Other Membrane  Protein Links •MemProtMD: A database of membrane proteins embedded in lipid bilayers with lipids obtained in Molecular Dynamics simulation iCn3D Basics: Saving Files File, Save File • In a new iCn3D window choose Open File,  iCn3D PNG image and see the same file you started with. • Likewise, in a new iCn3D window choose Open File, State/Script File and see the same file you started with. • They can be sent to others to open as well iCn3D Basics: Analysis Menu Analysis Menu iCn3D Basics: Mouse Commands iCn3D modeling screen Mouse commands rotate:  click and drag (mouse: left click and drag; keyboard: j, i, l, and m keys) zoom: pinch and spread (mouse: rotate the scroll wheel; keyboard: x and z keys) translate: two finger click and drag (mouse: right click and drag) Re-center: left click View from the top menu bar, then select “Center Selection” Note:  ctrl click on a PC = command click on Mac alt click on PC = option click on Mac iCn3D Basics: Selecting and Viewing with a mouse Selecting with the mouse (left) and viewing selection (right) iCn3D Basics: Style and Color Style and Color iCn3D Intro Tutorial A: Modeling a Short Peptide in a Protein A. Modeling short sections of a protein chain Pick one of the small protein fragments below for modeling using iCn3D PDB Description of protein (all small fragments) 2YW8 Crystal structure of human RUN and FYVE domain-containing protein 6EEY human Scribble PDZ4 R1110G Mutant 2PA1 PDZ domain of human PDLIM2 bound to a C-terminal extension from human beta-tropomyosin 3A03 Hox11L1 homeodomain 3IWL cisplatin bound to a human copper chaperone (monomer) 5Z2S DUX4-HD2 domain 6L1C PHF20L1 Tudor1 Y24L mutant 3D2N MBNL1 tandem zinc finger 1 and 2 domain 3RD2 NIP45 SUMO-like Domain 2 7NZC SH3 domain of POSH (Plenty of SH3 Domains protein) 1I2T HUMAN HYPERPLASTIC DISCS PROTEIN: AN ORTHOLOG OF THE C-TERMINAL DOMAIN OF POLY(A)-BINDING PROTEIN 1NTE CRYSTAL STRUCTURE ANALYSIS OF THE SECOND PDZ DOMAIN OF SYNTENI 2Y9U Structural basis of p63a SAM domain mutants involved in AEC syndrome 2FMA Alzheimer's Amyloid Precursor Protein (APP) Copper Binding Domain in 'small unit cell' form, atomic resolution 4OU0 Crystal Structure of RPA32C 1ZT3 C-terminal domain of Insulin-like Growth Factor Binding Protein-1 isolated from human amniotic fluid 2E3H Crystal structure of the CLIP-170 CAP-Gly domain 2 1L9L GRANULYSIN FROM HUMAN CYTOLYTIC T LYMPHOCYTES 5EFM Beclin 1 Flexible-helical Domian (FHD) (141-171) 2BZX Atomic model of CrkL-SH3C monomer 1NHL SNAP-23N Structure 7UW7 Crystal structure of the Human TRIP12 WWE domain (isoform 2) in complex with ADP 4N7F 3rd WW domain of human Nedd4-1 2Q9V C890S mutant of the 4th PDZ domain of human membrane-associated guanylate kinase 6T9Q second, C-terminal repeat of the DNA-binding domain of human TImeless 1WVN domain 3 of human alpha polyC binding protein 3I8Z human chromobox homolog 4 (CBX4) 2F60 Dihydrolipoamide Dehydrogenase (E3)-Binding Domain of Human E3-Binding Protein 7FGN FAF1 UBL1 5UM3 V122L mutant of human UBR-box domain from UBR2 2FJZ Alzheimer's Amyloid Precursor Protein (APP) copper-binding domain (residues 133 to 189) in 'small unit cell' form, metal-free 2.      Input in your assigned pdb code and select Load Biological Unit 3.      Choose Analysis, Seq and Annotation 4.       Choose Details tab and uncheck conserved domains 5.    With your mouse, select, hold, and sweep between the first 5-10 amino acids (given in single letter code) as illustrated below. When you select them, they will turn yellow. 6.      Choose View, View Selection (to limit view to what you want 7.       Choose Style, Proteins, Sticks to see all the bonds 8.       Change the background from black by choosing from top menu bar Style, Background, Transparent 9.      Choose, Analysis, Label, Per residue/#; then Analysis, Label Scale, 2 10.  Next, color your model as shown below in different ways as described in the table below.  Then take a screen capture of the selection and replace the image in the table cell with your own Color Paste snip of renderings as shown below. Spectrum, Selection to better see each amino acids in selection Charge Gray if no charges (ignore yellow highlight) Hydrophobicity (if nonpolar like oil) Atom (red oxygen, blue nitrogen, yellow Sulfur

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/iCn3D_Molecular_Modeling_Tutorials/iCn3D_Basics%3A_File_Types.txt

B.  Rendering Full Proteins Pick one of the proteins below that has both alpha helices and beta sheets.  You will then  change the protein style (rendering) to see the same protein in different ways to illustrate different properties of the proteins. Monomeric proteins with alpha (helices) and beta (sheets) 1HDO Human biliverdin IX beta reductase: NADP complex 2HC2 Engineered protein tyrosine phosphatase beta catalytic domain 5ZUN Crystal structure of human monoacylglycerol lipase in complex with compound 3l 1X3S Crystal structure of human Rab18 in complex with Gppnhp 4IN0 Crystal Structure of human splicing factor dim2/TXNL4B 1KGD Crystal Structure of the Guanylate Kinase-like Domain of Human CASK 5KQL Co-crystal structure of LMW-PTP in complex with 2-oxo-1-phenyl-2-(phenylamino)ethanesulfonic acid 1QGV HUMAN SPLICEOSOMAL PROTEIN U5-15KD 1MF7 INTEGRIN ALPHA M I DOMAIN 4RQR Crystal Structure of Human Glutaredoxin with MESNA 4JKA Open and closed forms of R1865A human PRP8 RNase H-like domain with bound Co ion 5C4M RhoA GDP with novel switch II conformation 6P0J Crystal structure of GDP-bound human RalA 4MMM Human Pdrx5 complex with a ligand BP7 4M6IJ Crystal structure of human dihydrofolate reductase (DHFR) bound to NADPH 3M9J Crystal structure of human thioredoxin C69/73S double mutant, reduced form 1.       Load ID  4LPK, Crystal Structure of K-Ras protein with a small molecule, GDP, bound 2.      Choose Color, Secondary, Sheet in Yellow 3.      Style, Background, White 4.      Choose Style, Proteins and display as ribbon, cylinder and plate, C alpha trace, backbone, lines and sphere 5.      Paste your results in the table below. 6. Results (replace image with yours) PDB ID, description: ribbon Cylinder and plate C alpha trace backbone line sphere 7.       Model a protein dimer that has two subunits.  Use the pdb code for 1LFD (CRYSTAL STRUCTURE OF THE ACTIVE RAS PROTEIN COMPLEXED WITH THE RAS-INTERACTING DOMAIN OF RALGDS). Paste your favorite image below. 8.      Model a huge structure, the human rhinovirus 14 (causes colds, PDB: 4RHV). It so big you have to load it in a different way, as shown below. Paste your favorite image below. iCn3D Intro Tutorial C: Finding Pockets in Proteins Small molecules that bind to larger proteins must have shape AND charge complementarity with the binding pocket in the protein.  You can put a small molecule into an appropriate-sized pocket in a protein.  You can’t put a positively charged small molecule into a pocket lined with a positive charge.  Let’s find the pockets in a small protein, LMWPTP, a phosphatase that cleaves a negatively charged phosphate group (PO3-2) proteins (pdb 1xww).  It also binds the small sulfate ion (SO4-2) in the same pocket. Finding Pockets Let’s find the pocket where the ligand could bind using a free program called CavityPlus. 2022 1.      Load http://www.pkumdl.cn:8000/cavityplus/computation.php#/ and select Start Computing 2.      Input 1xww, then select Click to Search. Wait until the structure loads to continue. 3.      Then simply choose Submit.  (Make sure that Use Ligand Mode is not selected) bnbnbn 4.      After the run, you will see a new window open on the left-hand-side with the protein and the top #1 Cavity  highlighted.  Site 1 is the presumptive location for the binding of SO42-. Use your mouse to rotate the protein to better see the cavity.  To see a list of the amino acids lining the binding pocket surface, and the surface area and volume of the cavity, select under More. They will appear in the Residue row. 5.  Copy and Paste into the table below the list of amino acids comprising the pocket into the table below. Then take a screen snip as shown in the image to the right (Note:  if you can unzip the downloaded file, you could select the Download Results link and use other programs to view the results). Amino acids in pocket Image snip Viewing  Small Molecule in a Binding Pocket Now let’s model the phosphate (PO3-2)/sulfate (SO4-2) binding site in the phosphatase using iCn3D. 1.  load 1XWW in iCn3D 2.  Render the proteins as follows: • Analyses, Sequences and annotation, Details Tab, uncheck conserved domain • Click 1XWW_A 1st, then in the top menu bar choose Select, Save Selection and name it phosphatase • Color, Charge • Style, Surface Opacity, Fast Transparency, 3 • Style, surface type, molecular Surface 3.  Next render the SO4 as follows: • Choose SO4 (2) with the mouse, then Select, Save Selection and name it SO4 • Style, Chemicals, Sphere 4.  Snip and paste an image of the protein with the surface display and the bound SO4-2 in spheres. 5.  Optional: To see actual interactions between SO4-2 and the protein • Style, Remove surface • Analysis, Interactions, •  In popup window, choose for 1st set – sulfate; choose for the 2nd set – phosphatase; click 3D Display Interactions; Snip table with types/colors on interactions • View Selection; Style, Sidechains, Stick; Color, by Atom, • Analysis, Label, Per Residue and number; Analysis, Label Scale, 2; • Style, background, white • Snip an image of the interaction legend and modeled interactions, and paste below.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/iCn3D_Molecular_Modeling_Tutorials/iCn3D_Intro_Tutorial_B%3A_Rendering_a_Protein.txt

Use iCn3D to model the binding of one of the psychoactive/analgesic drug to their receptor (either the 5HT or cannabinoid receptor). Paste the final model in the space shown. Serotonin (5-hydroxytryptamine) receptors 5-HTreceptors are indirectly involved in the mechanism of action of antidepressant drugs. Most antidepressant drugs like Prozac increase the concentration of 5-HT in the extracellular brain synapse by inhibiting its reuptake into neurons.   Prozac doesn’t bind to the 5HT receptor but to a membrane protein that removes 5HT from synaptic region. There are 7 types of 5HT receptors and each has different biochemical effects Different drugs target different 5HT receptors. Let’s focus drugs that target 5-HT2A act as antidepressants,  but also lead to hallucination. Serotonin (5HT) 2A receptor:  These molecules bind to it. • serotonin (5HT), the physiological agonist, 7WC4 (does not cause hallucinations) • psilocin, 7WC5 , hallucinogenic, a metabolite of psilocibin) • LSD, 7WC6, hallucinogenic • lisuride, 7WC7, non-hallucinogenic • IHCH-7086, nonhallucinogenic THC and CBD receptors These molecules bind to them. • Human CB1 in complex with agonist AM11542 (5XRA) • class A GPCR Cannabinoid Receptor-Gi Complex Structure with bound agonist (6KPF) - The agonist is AM12033, which is similar to AM11542 •  CBD-bound full-length rat TRPV2 in nanodiscs (6U88) Your model: PDB ID: "DRUG” RECEPTOR: Rendered Image iCn3D Skill: Alternative Rendering and Saving Files Structure • PDB ID: 1xww • Protein:  Low molecular weight protein tyrosine phosphatase • Activity:  hydrolyzes Tyr-OPO32-  phosphoester bond • Description: single chain, bound SO42- (competitive inhibitor), bound glycerol (nonspecific stabilizer) Alternative Rendering Load Structure and Mouse/Trackpad Controls • Open iCn3D - https://www.ncbi.nlm.nih.gov/Structure/icn3d/full.html • For a simple menu, use the dropdown: File > Customize Menus > Simple Menus. • In the Please input MMDB or PDB, enter 1xww. Press enter or click load biological unit. • Default render is ribbon (cartoon) with black background and small molecules shown as sticks. Hover over objects with the mouse to reveal their identity. 1. From the top menu bar, choose Style, Protein, then try some of the available choices: 1. For your favorite protein styles, select Color by left-clicking on the top menu, then pick available choices. Try: • Secondary, Sheets in Yellow • Charge • Hydrophobicity 1. Under Style, choose ProteinRibbon. Under Color, choose Secondary, Sheets in Yellow before the next step. 2. To view sidechains, Style, Side Chains, Stick (they will remain the same color as secondary structure for now) 3. Color for SO42- and bound glycerol will default to CPK coloring (key below) 4. Convert back to cartoon (Style, Side Chains, Hide) Saving Files 1. Style, Background, Transparent 2. Saving Files: There are several ways to save your work. The first option below saves a PNG image, the second creates a share link 1. File, Save Files, iCn3D PNG image, original size;  Give it a name. Can be reloaded in iCn3D with File, Load, iCn3D PNG IMAGE 2. File, Share Link,  Save Lifelong Short URL. Copy and paste this link to share your work. Pre-Rendered Model Link To check your work (or if you got stuck during any of the steps above) catch up using this link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?CgfEnF27TN7aYQpr6 iCn3D Skill: Displays surface of a protein - Superoxide Dismutase A.  The molecular surface of superoxide dismutase PDB ID: 2sod • Description • superoxide dismutase • dimer with 2 Cu2+ ions bound in each subunit • catalyses the reaction of O2- + O2- + 2H+ →  H2O2 + O2 Instructions 1. File, Retrieve by ID, MMDB 2. Style, Surface Opacity, Fast Transparency, 0.2 3. Select, Select on 3D, Chain.  Alt Click on one chain 4. Color, Secondary, Sheets in Yellow 5. Style, Surface Type, Molecular (takes a while to render) 6. Now Alt Click on the other chain 7. Color, Wimley White Hydrophobicity(takes a while to render) 8. Style, Surface Type, Molecular (takes a while to render) 9. Style, Background, White 10. File, Share Link, Save short url B.  The electrostatic surface potential of superoxide dismutase Background:  Superoxide (O2-) is a toxic free radical and hence dangerous.  To capture it as effectively as possible, the surface distribution of charged side changes is such that a positve electrostatic potential surrounds each active site. 1. File, Retrieve by ID, MMDB 2. Select, Select on 3D, Chain.  Alt Click on one chain, Control-click on the second.  Alternatively, choose Analysis, Sequence and Annotations, Details Tabs (uncheck conserved domains), and click one chain (blue font) and then control-click the second so both are highlighted. 3. In the Details tab of Analysis, Sequence and Annotations, sweep both Cu ions (with mouse) to select them, the Color, Yellow 4. Analysis, DelPhi Potential, DelPhi Potential 5. Choose the Surface with Potential Map tab.  Chose the options below 6. Select Surface with Potential.  Look a the blue (positive potential) surround the yellow Zn ions in both active sites. 7. Style, Surface Type, Molecular (takes a while to render) 8. Style, Background, White 9. File, Share Link, Save short url

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/iCn3D_Molecular_Modeling_Tutorials/iCn3D_Intro_Tutorial_D%3A__Modeling_Psychoactive_Drugs_in_Target_Proteins.txt

Superimpose and annotate active (phosphorylated) form of cyclin-dependent kinase 2 (1JST) and inactive (1FIN) Description 1JST (active) • PHOSPHORYLATED CYCLIN-DEPENDENT KINASE-2 BOUND TO CYCLIN A.  Has bound Mn2+ and ATPγS. • pT160 is on the regulatory T-loop of CDK2.  It is mostly buried and neutralized by 3 Arg side chains. Compared to unphosphorylated CDK2–CyclinA, the T-loop moves by as much as 7 Å, • T loop about  147-163 1FIN (inactive): • CYCLIN A-CYCLIN-DEPENDENT KINASE 2 COMPLEX.  Has a bound ATP as well as Cyclin A (B and D chains) Instructions 1. File, Align, Structure to Structure 2. Input 1JST and 1FIN 3. Choose All Matching Molecules Superimposed 4. In the Select residues in aligned alignment window, click  the blue font 1JST-C to select the entire chain.  Color, Green, Dark Green (green for go = active) 5. Likewise, select 1FIN-A.  Color, Yellow, Yellow (so we can color the T loop red) 6. Alt Click in the modeling window MN and ctrl Click ATP.  Select, Save Selection, MN_ATP 7. Style, Chemicals, Sphere 8. In long sequences, capture with mouse approx T loop 147-163. Select, Save Selection, T loops; Color red 9. In the long sequences, hover over and then click to select in 1JST_C  x160 (this is phospho-Thr160) and then in 1FIN_A  T160 (i.e. Thr 160). Select, Save Selection, Thr160s 10. Style, Sidechains, Sticks 11. Color, Atom 12. In the Defined Sets, click then ctrl click all of the selections you made: MnATP, Thr160s, align_1FIN_A, and align_1JST_C.  Then save as 000All. Using 000 at the start of the name put the combined selection at the top of the Defined Sets window.  This will be important later. 13. Make sure 000All is selected, then View,  View Selection 14. Style, Background, White 15. File, Share Link, Lifelong Short Link, 16. Paste the link into a new window, much as a student would.  Analysis, Defined Set, then click 000All at the top to highlight the superimposed chains.  Once all is highlighted, toggle between the two states by clicking the Alternative (Key “a”) menu button directly under the File button. 17. Now try View, Side by Side to see both (don’t toggle this form) iCn3D Skill: Analysis of Noncovalent Interactions Structure • PDB ID: 3K83 • Protein:  Inhibitor (below, abbreviated F278458) bound to a humanized variant of Fatty Acid Amide Hydrolase (FAH) • Activity:  FAH Catalyzes the hydrolysis of endogenous amidated lipids like the sleep-inducing lipid oleamide, the endocannabinoid anandamide, and other fatty amides, regulating the signaling functions of these molecules • Description:  The functional unit is a dimer, each with an active site.  A chloride binds between the two subunits In this activity you will see key noncovalent interactions between the inhibitor and (FAH).   You will pick one inhibitor and view its noncovalent interactions with one subunit Modeling Instructions 1. Open a new iCn3D window, 3K83 at: https://www.ncbi.nlm.nih.gov/Structure/icn3d/full.html 2. Select, Select on 3D to ensure default is Residue; With mouse, zoom and center to Alt-Click the inhibitor (F278458) in the magenta subunit (option-click on a Mac). 3. Select, Save Selection, name it Drug 4. Analysis, Interactions. 5. Under 1. Choose interaction types and their thresholds, Check only the noncovalent interactions shown below. Make sure the Contacts/Interactions is unchecked as it shows all interactions between contacting Van der Waals surfaces including many hydrophobic interactions, so it is quite cluttered. 1. Select the Drug as the first set (under 2. Select the first set); choose 3K83 (under 3. Select the second set) 1. Click on the box: 4. 3D Display Interactions. This will display interacting side chains.  Drag the HBonds/Interactions window to the bottom right away from the molecular display. Note the coloring of the dotted lines: Green - hydrogen bonds Red - π-cation Blue - π stacking 1. Select, Save Selection, name it Interactions 2. Style, Sidechains, Stick (so the next step will color side chains) 3. Color, atom (Note: this drug is covalently bound to Serine241) 4. In defined sets Ctrl click Drug and Interaction to highlight all 5. View, View Selection (to only see Drug and Interactions) 6. Select, Toggle Highlight (to remove highlights if necessary) Note: Once you run interactions, iCn3D adds many new additions to the Defined sets window.  Explore these to learn different ways to examine the interactions. Optional (if you’d like to add labels, recolor the background, and obtain the share link as in the previous activity) 1. Analysis, Label, Per Residue & No 2. Analysis, Label Scale, pick a number that works for you 3. Style, Background, Transparent 4. File, Share Link, copy short link Pre-Rendered Model Link To check your work (or if you got stuck during any of the steps above) view the model using this link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?pDr5EBZmo3TyAbTP6 Note: You can alter interactions on the 3D model, or examine 2D displays of them. Try: ●     Click 5. Reset at the bottom of the HBonds/Interactions window. Add Contacts/ Interactions (mostly nonpolar) by clicking on the box. Choose Drug as the first set. Choose 3K83 as the second set. Then click 4. 3D Display Interactions to show. ●     Next, try 4. 2D Interaction Network. In the popup that opens, click a colored line in the 2D window to highlight a specific interaction. If using the Pre-Rendered model: From the top menu, select: AnalysisInteractions. In the popup choose 5. Reset, select 3K83 (full protein) as set 1 and drug as set 2, then 4. 2D Interaction Network. Click colored lines in the 2D window.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/iCn3D_Molecular_Modeling_Tutorials/iCn3D_Skill%3A__Aligning_two_structures.txt

Structure • PDB ID: 3UBB • Protein:  Rhomboid intramembrane serine protease GlpG (3UBB) with phosphonofluoridate inhibitor • Activity:  Integral membrane serine protease • Description:  Single chain transmembrane protease from E. coli bound to a phosphonofluoridate inhibitor, which is covalently bonded to the catalytic serine. Red and blue dots (“dummy” atoms) indicate extracellular and intracellular membranes, respectively. Uses a catalytic dyad composed of serine (S201) and histidine (H254). Load Structure 1. Open iCn3D - https://www.ncbi.nlm.nih.gov/Structure/icn3d/full.html 2. In the Please input MMDB or PDB, enter 3UBB.  Press enter or click load biological unit. Selecting using: 1. The structure viewer window with the Mouse and 2. The sequences and annotations menu (Key Learning Objectives) This section will show how you can select residues, chains, etc in the Modeling window. 1. Select, Select on 3D to ensure default is Residue; With cursor, hover over the inhibitor (name 3UB) and Alt Click (option click on Mac) it.  A yellow halo will appear around it. 2. Select, Save Selection, name it Inhibitor. 3. Now, use the top menu to open the sequences and annotations tab: Analysis, Seq. and Annotations 4. In the sequences and annotations window, uncheck “Conserved Domains,” and then click the Details tab. Click individually on the one letter code for S201 and H254. On selection, they will turn yellow. (Note: To see the scroll bar in sequences/annotations in a Mac, choose Systems preferences in your operating system [not the iCn3D settings], General, show scroll bars and check always; see iCn3D About notes). 1. Select, Save Selection, name CatDyad Rendering (Optional, can use the highlighted link below instead) 1. Select, defined sets (Note that “defined sets” brings up a complete list of objects, many selections are pre-built into iCn3D to get you started with a model.) 2. In Selected Sets, click 3UBB_A. 3. Color, Unicolor, Gray, Light Gray 4. In Selected Sets, click CatDyad 5. We want to show the side chain of the catalytic dyad, so use the top menu: Style, SideChains, Sticks 6. Recolor to CPK coloring: Color, Atom 7. Analysis, Label, Per Residue and Number 8. Analysis, Label Scale, pick number that works for you 9. Style, Background, Transparent 10. Remove any active selections (yellow glow) by Select, Toggle highlight Pre-Rendered Model Link https://structure.ncbi.nlm.nih.gov/icn3d/share.html?fhT3dwckYg8i5XJj8 The short URL above may be used to catch up for the next section of the tutorial Selecting sphere within around 5Å of the inhibitor (Key Learning Objective) Our goal is to find all atoms with 5Å from the inhibitor, this time without showing interactions. In the next step, we will designate 2 sets of objects. Set 1 will be the inhibitor. Set 2 will be nearby residues/bound molecules. In our case, Set 2 will be defined as the protein. 1. Select, by Distance 2. For the first set, select inhibitor; for Set 2 click 3UBB 3. For Set 2. Sphere with a radius to 5 Å 4. Click Display; Close the Select by distance window 5. Now save the highlighted groups 5Å from the inhibitor through Select, Save Selection, Name 5AfromInhib 6. Style, Side chains, Stick 7. Color, Atom 8. Select both the inhibitor and the surrounding residues: In Defined Sets, Click Inhibitor, and Ctrl+Click 5AfromInhib (Command+Click on a Mac) 9. Display only the active site for clarity: View, View Selection 10. Analysis, Label, Per Residue & Number 11. To see water, Style, Water, Sphere 12. File, Share Link, copy short link Pre-Rendered Model Link To check your work (or if you got stuck during any of the steps above) view the model using this link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?sqZf4Zvqn5zWK21d9 This link shows the membrane. Choose View, Toggle Membrane to hide it.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/iCn3D_Molecular_Modeling_Tutorials/iCn3D_Skill%3A__Creating_and_Saving_Selections.txt

Analysis: Mutation A simple way to change 1 amino acid to another PDB ID: 1xww Description:  human B-form low molecular weight protein tyrosine phosphatase (has single domain) with bound sulfate (SO4) and glycerol (GOL) Instructions 1. File, Retrieve by ID, MMDB 2. Analysis, Mutation 3. Input the desired mutation C12S this way (1XWW_A_12_S)  where A is the A chain, 12 is the aa # and S is the new mutated amino acid residue Ser. Note that C12 is the wild-type active site nucleophile in phosphatases and changing it to Ser will effectively abolish phosphatase activity.  To view you could use 3 different methods.  We'll use one. 4. 3D with scap:  toggle back and forth between structures using the key “a” 5. Interaction:  shows the mutation in 3D and change in interactions 6. PDB:  shows structure and exports a PDB file within 10 A of the mutation 7. Choose Interactions and say wow! 8. Color, Atom 9. Style, Background, White 10. Toggle back and forth between the structures with the letter “a” 11. File, Share Link, Lifelong Short Link iCn3D Skill: Selection through Sequence and Annotations Structure • MMDB ID: 1xww • Protein:  Low molecular weight protein tyrosine phosphatase • Activity:  hydrolyzes Tyr-OPO32-  phosphoester bond • Description: single chain, bound SO42- (competitive inhibitor), bound glycerol (nonspecific stabilizer) Load Structure and Mouse/Trackpad Controls • Open iCn3D - https://www.ncbi.nlm.nih.gov/Structure/icn3d/full.html • For a simple menu, use the dropdown: File > Customize Menus > Simple Menus. • In the Please input MMDB or PDB, enter 1xww. Press enter or click load biological unit. • Default render is ribbon (cartoon) with black background and small molecules shown as sticks. Hover over objects with the mouse to reveal their identity. Figure: The Sequences and Annotations Menu For this part, we will be using the model from Activity 1. To load the premade model from Part 1, use this link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?CgfEnF27TN7aYQpr6 From the literature, it is known that the active site is a nucleophilic cysteine (C12).  It is part of the phosphate-binding loop (P-loop, AA 12-18: sequence CLGNICR). Let’s find, select, and render these amino acids. Modeling Instructions 1. Under Analysis (top menu bar), choose Sequence and Annotations 2. Choose Details tab, uncheck Conserved Domains Before we continue, look at the built-in choices you have for selection: 1. In the Sequences and Annotation window, click Protein 1XWW_A 2. Under Select (top menu bar), choose Toggle Highlights 3. Hover over C12 in the sequence (in Seq and Annot window), click and hold down the mouse key, and sweep over C12-C18 to select the P loop 4. Select, Save Selection, name it: Ploop 5. Within this highlighted selection, Style, Side Chains, Sticks 6. Color, Atom 7. Analysis, Label, Per Residue & Number 8. Analysis, Label Scale, pick number that works for you 9. Analysis, Label, Change Label Color (globally). Click in the text box and a Color box will pop up,  choose from a palette, then Display.  (Alternatively, pick a hex code). 10. In the Sequences and Annotation window, click SO4 11. Style, Chemicals, Sphere to change the sulfate to a space filling rendering 12. Files, Share Link, Copy Short URL Pre-Rendered Model Link To check your work (or if you got stuck during any of the steps above) catch up using this link: https://structure.ncbi.nlm.nih.gov/icn3d/share.html?QhtGuE8pkaJpGs1X9 Note: For some enzymes, iCn3D can automatically display key active site and binding residues. These can be seen as shown by selecting the items indicated in the left figure.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/iCn3D_Molecular_Modeling_Tutorials/iCn3D_Skill%3A__Mutations.txt

Protein:Protein Interface PDB ID: 3S9D • binary complex between interferon alpha 2 (IFNa2) and its recetor IFNAR2 • Cyan:  IFN, blue IFN receptor Instructions 1. Retrieve by ID, MMDB 2. Analysis, Seq. & Annotationsm, Details Tab. 3. Select magenta IFN alpha chain:  3s9d_A 4. Select,  Save Selections INF 5. Repeat with blue 3s9d-b. Save Select INFR 6. Select, INF; Select by Distance, within 5A.  Set 1: selected;  Set 2: nonselected; Display 7. Select, Save Selection:   5Ang from IFN 8. Select 5A from INF:  Style, side chains stick, color atom 9. repeat for INFR 10. Select Select by Distance, within 5A. Set 1: IFNR; Set 2: nonselected;  Display 11. Select, Save Selection:   5Ang from IFNR 12. Select, Side Chains, sticks; Colore atom 13. Select 5A from IFNR: Style, side chain sticks.  Color, charge, Style, surface type, vanderWaals 14. Analysis, Interactions, Set 1: 5A from INF; Set 2: 5A from INFR 15. Select, Save, Selections, Inteactions 16. Style, Background, White 17. File, Share Link, Lifelong iCn3D Tutorial: Binding interactions of SARS-Cov-2 Spike receptor domain A.  1st Way PDB ID: 6M0J Description:  SARS-CoV-2 spike receptor-binding domain bound with ACE2 • Angiotensin-converting enzyme 2, chain A, denoted 6M0J_A, pink • Spike Protein S1, chain B, denoted 6M0J_E, blue Instructions 1. File, Retrieve by ID, MMDB 2. Seq. & Annotations, Uncheck Conserved Domains 3. Analysis, Interactions, then select the choices in the image below.  (Leave out Contact/Interactions to decrease clutter) 4. Click 3D Display Interactions 5. Without closing Interactions window, choose Select, Save Selection, name it BindingInterface 6. Click 2D interaction network to get the image below 7. In the Selected Sets window, choose BindingInterface. 8. View, View Selection 9. Select, Select Side Chain 10. Color, Atom 11. In the Selected Sets window, choose BindingInterface. 12. Analysis, Label, Per Residue & Number 13. Style, Background, White 14. File, Share Link, Lifelong Short Link B.  2nd Way Instructions 1. File, Retrieve by ID, MMDB 2. Seq. & Annotations, Uncheck Conserved Domains, Details Tab 3. Check Interactions 4. From right hand side window sequences: Under 6M0J_A, select the blue label Interact .E,  Save as A_with_E 5. Under 6M0J_E, select the blue label Interact .A,  Save as E_with_A 6. Analysis, Interactions. 7. Choose A_with_E and E_with_A 8. 3D Display Interactions 9. Select, Sidechains 10. Color, Atom 11. In H Bonds/ Interactions window, choose Highlight Interactions in Table (can toggle on/off individual interactions), 12. Try the 2D Interaction Map and the Buried Surface Area 13. Style, Background, White 14. File, Share Link, Lifelong Short Link iCn3D Tutorial: Overlay many ACE2 receptor binding domain analogous to SA Use BlastP to align receptor-binding domains PDB ID: 6M0J • SARS-CoV-2 spike receptor-binding domain bound with ACE2 • Angiotensin-converting enzyme 2, chain A, denoted 6M0J_A, pink • Spike Protein S1, chain B, denoted 6M0J_E, blue Instructions 1. File, Retrieve by ID, MMDB 2. Seq. & Annotations. In Summary tab check Custom and Conserved Domains 3. In Summary  tab, click on 6M0J_E 4. View, View Selection 5. In Details tab, Add track. 6. Choose the FASTA Alignment Tab in the new window 7. Cut and paste the aligned FASTA sequences file you made using the instructions on the next page or to save time just copy/paste the same FASTA Alignment  file below into the iCn3D Fasta window. 8. Enter 319 for the Position of the first residue in Sequences & Annotations window (# is right there);  Check Color sequence by identity: 9. Choose Add Tracks.  The RBD appears color coded with the darkest rd 100% identity and the darkest blue 0% (See F377, K387) 10. Style, Background, White 11. File, Share Link, Lifelong Short Link (this might not work if you choose too many proteins) Instructions to use Blast Use of BlastP for COV-2 ACE2 Receptor Binding Domain (RBD) Find sequences homologous to the structure of the RBSthrough BlastP: Instructions - use this image for help 1. Go to BlastP 2. Enter this Accession number (in this case PDB ID with chain:  6M0J_E for the RBD 3. Under Choose Search Set 4. Database:  Choose UniProt 5. Organism:  input Coronavirus and click it when it appears in the search 6. Add Organism and then input SARS-CoV-2 and click it when it appears in the search;  Click Exclude (since we already know the structure of the RBD of CoV-2) 7. Choose Blastp and then BLAST 8. After a few minutes you will get the general output shown below . 9. Check just the ones with percent identity >30% and choose Download, Fasta Aligned  Sequences 10. Copy the resulting text file and use in iCn3D as described in the previous page.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/iCn3D_Molecular_Modeling_Tutorials/iCn3D_Skill%3A__Showing_a_Protein-Protein_Interface.txt

Protein Kinase B (AKT) A more complicated protein with multiple domains and at least two major conformations with different PDB structures: Protein Kinase B (aka AKT) PDB files: • 3CQW (active) • 3O96  (inactive) • 1UNQ  PH domain A.  Exploration of AKT – Domain Structure • PDB ID: 1UNQ • Description: Pleckstrin Homology Domain Of Protein Kinase B/Akt Bound To Ins(1,3,4,5)-Tetrakisphophate 1. Open new iCn3D 2. File, Retrieve by ID, MMDB 3. Alt Click the ligand.  Style, Chemicals, Sphere 4. Select, Select on 3D, check Chain, alt click the protein 5. Color,  Secondary, Sheets in Yellow With the protein still selected: 1. Analysis, Interactions 2. For the second set choose chemicals 3. Check just the noncovalent interactions shown below 4. Choose 3D Display Interactions.  This will display interacting side chains 5. Selections, Save Selections, name it Interactions 6. Style, Sidechains, Stick (so next step will just color side chains) 7. Color, atom 8. In defined sets choose Interactions.  Analysis, Label, Per Residue & No. 9. Analysis, Label Scale, 3 10. Style, Background, White 11. File, Share Link, copy short link B.  Exploration of AKT – Self-defined aa, sequences, Render PH-domain, N-lobe, C-Lobe, Catalytic Loop, Activation Loop PDB ID: 3O96 (inactive) Description • Crystal Structure of Human AKT1 with an Allosteric Inhibitor • RAC-alpha serine/threonine-protein kinase; Chain A • Allosteric inhibitor IQO Cartoon structure Instructions 1. File, Retrieve by ID, MMDB 2. Analysis, Seq. & Annotations, Uncheck Conserved Domains 3. Analysis, Defined Sets.  Click 1st domain (PH = top 3O96_A_3d_domain) 4. Style, Protein, C-Alpha Trace. 5. Color , Unicolor, Yellow 6. Defined Sets.  Choose 2st domain (N-Lobe).  Color, unicolor, Cyan 7. Defined Sets.  Choose 3st domain (C-Lobe).  Color, unicolor, Grey, Light Gray 8. Seq. & Annotations, Details  Tab, Uncheck Conserved Domains. 9. Select by hand the catalytic  loop  (table below) by using the horizontal scroll to find and select 271V-287H.  Hover over each amino acids and click to select it or use your mouse to sweep out a rectangle over all the range of amino acids. 10. Select, Save Selection, CatLoop 11. Color, Unicolor, Red 12. Repeat for Activation Loop and save the selection. Color Unicolor, Blue 13. Style, Background, White 14. File, Share Link, Lifelong Short Link Numbering or generic kinase and AKT SITE Generic Kinase Akt (+108) N lobe K72 K180 N lobe E91 E199 C lobe, cat loop R165 R273 C lobe, cat loop D166 D274 C lobe, act loop D184 D292 C lobe, act loop T197 T305 (NO SEQ) Approx Cat Loop 163-179 271V—287H Approx Act loop Start DFG (292-294) to APE 184-200 292-319 308Tmiss toAPE end 319 C.  Compare 3CQW (active) and 3O96  (inactive) – Superimpose two AKT structures PDB ID: 3CQW and 3O96 Description 3O96 (inactive): Crystal Structure of Human AKT1 with an Allosteric Inhibitor • RAC-alpha serine/threonine-protein kinase; Chain A • Allosteric inhibitor IQO 3CQW (active): • Crystal Structure of Akt-1 complexed with substrate peptide (hence active form) and inhibitor • Peptide:  Glycogen synthase kinase-3 beta peptide Instructions 1. File, Align, Structure to Structure 2. Input 3CQW and 3O96 3. Choose All Matching Molecules Superimposed 4. In the top of the right hand alignment window, save the aligned state as presented (al_seq1).  Just the aligned sequences will be highlighted. 5. View,  View Selection 6. Style, Background, White 7. File, Share Link, Lifelong Short Link 8. Toggle between the two states by clicking the toggle menu button just underneath the File menu button iCn3D Tutorial Question: Intermediate Problem - Cyclooxygenase II Structure • PDB ID: 4PH9 • Protein:  Ibuprofen (IBP) bound to cyclooxygenase-2 • Activity:  COX-2 catalyzes the conversion of arachidonic acid(AA) to prostaglandin G2 (PGG2), and is a target of non-steroidal anti-inflammatory drugs (NSAIDs) and COX-2 selective inhibitors (coxibs).  Arachidonic acid (AA), not shown in this structure, binds in a “L” shape Key amino acids • Arg-121 and Tyr-356 are close to the carboxylate of AA • Phe205, Phe209, Val228, Val344, Phe381, and Leu534 form a hydrophobic groove for the ω-end of AA. • Ser 530, which is above this, gets acetylated by aspirin • Tyr 385, near C13 in AA, forms a free radical which removes a single electron from C13 For more information on the mechanism of COX-2, scroll down to the end of the chapter section in Fundamentals of Biochemistry. Description:  Biological dimer with heme, ibuprofen (IBP), and many other ligands bound. The initial display is messy!  Your task is to clearly render a specific structural feature. Assessment: The enzyme cyclooxygenase-2 (COX-2) produces arachidonic acid from prostaglandin G2, as shown in the reaction below. Prostaglandin G2 is an important metabolite in inflammation, so inhibition of (COX-2) reduces inflammation. Potential visualization activities for students: 1.  Identify the noncovalent interactions of COX-2 with a heme in 1 subunit 2. Model and describe the noncovalent interactions of COX-2 with ibuprofen 3. Model the interactions at the dimer interface of the protein; identify two amino acids on different subunits that are participating in a [hydrogen bond / ionic interaction] 4. Show the key active site residues of the enzyme Steps for Discussion and/or Modeling STEP 1: As a table, or small group at one table, choose one from A-D above. STEP 2: Load the model 4PH9 at https://www.ncbi.nlm.nih.gov/Structure/icn3d/full.html With your group, broadly discuss the steps students would need to perform to accomplish the activity you chose in STEP 1. Consider: • for your course, would it make sense for them to start with the model as it is loaded? • If you were to pre-render the model and provide them a shared link, which steps would you perform ahead of time? Which steps of modeling are important to their learning STEP 3: Open the sequences and annotations tab: • Analysis → Sequences & Annotations • Uncheck “conserved domains,” and then check “functional sites” STEP 4: Discuss how the built in iCn3D features may help you create a model quickly. Pre-Rendered Model Links 1. Identify the noncovalent interactions of COX-2 with a heme in 1 subunit 1. Model and describe the noncovalent interactions of COX-2 with ibuprofen 1. Model the interactions at the dimer interface of the protein; identify two amino acids on different subunits that are participating in a [hydrogen bond / ionic interaction] 1. Show the key active site residues of the enzyme

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/iCn3D_Molecular_Modeling_Tutorials/iCn3D_Tutorial%3A__Protein_Kinase_B_%28AKT%29.txt

BIOMOLVIZ Promoting Molecular Visualization Literacy The BMV framework is used with permission from BioMolViz.Org Copy the appropriate row when assigning a theme, goal, and objective to a designated iCn3D or other biomolecular visualization assessment Atomic Geometry (AG) Three‐atom and four‐atom (dihedral) angles, metal size and metal‐ligand geometries, steric clashes AG1. Students can describe the ideal geometry for a given atom within a molecule and deviations from the ideal geometry due to neighboring interactions. AG1.01 Students can identify atomic geometry/hybridization for a given atom. (Novice) AG1.02 Students can measure bond angles for a given atom. (Novice) AG1.03 Students can identify deviations from the ideal bond angles. (Amateur) AG1.04 Students can explain deviations from the ideal bond angles due to local effects. (Amateur, Expert) AG1.05 Students can predict the effect of deviations from ideal bond angles on the structure and function of a macromolecule. (Expert) AG1.06 Students can identify the geometric features of bonds (e.g., peptide bond, glycosidic, phosphoester). AG2. Students can compare and contrast different structural conformations with regard to energy, the addition of substituents, and the impact on the structure/function of a macromolecule. AG2.01 Students can describe different conformations that a structure can adopt using visualization tools. (Amateur) AG2.02 Students can describe different conformations of atoms about a bond using visualization tools. (Novice) AG2.03 Students can distinguish energetically favorable and unfavorable conformations that a structure can adopt. (Amateur) AG2.04 Students can predict the effect of a given substituent on the structure and function of a macromolecule (e.g., substituent on a carbohydrate/ligand, R groups/rotamers, phosphorylation, methylation of nucleic acids, post-translational modifications). (Expert) AG3. Students can describe dihedral/torsion angles in biomolecules. AG3.01 Students can identify a dihedral/torsion angle in a three-dimensional representation of a molecule. (Novice) AG3.02 Students can identify the planes between which a dihedral/torsion angle exists within a three-dimensional representation of a macromolecule. (Novice) AG3.03 Students can identify phi, psi, and omega torsion/dihedral angles in a three-dimensional representation of a protein. (Amateur) Alternate Renderings (AR) Rendering of a macromolecular structure such as a protein or nucleic acid structure in various ways from the simplest possible way (connections between alpha carbons) to illustration of secondary structure (ribbons) to surface rendering and space filling. AR1. Students can interpret or create molecular images that convey features such as secondary structure, CPK coloring, and active sites. AR1.01 Students can manipulate rendered structures to illustrate molecular properties. (Novice) AR1.02 REMOVED (integrated with SF2.02) AR1.03 Students can describe or label structural differences among multiple structures. (Amateur, Expert) AR1.04 Students can infer information from rendering a structure in different ways. (Novice, Amateur, Expert) AR1.05 Students can create renderings that distinguish secondary structural features. (Novice) AR1.06 Students can create an information rich rendering of a structure that depicts structural features found in the literature. (Amateur) AR1.07 Students can create an information rich rendering of a structure containing ligands, covalent modifications, and noncanonical amino acids or nucleotides. (Amateur, Expert) AR1.08 Students can use molecular visualization to tell a story about a macromolecular structure. (Expert) AR1.09 REMOVED (integrated with MI1.02) AR1.10 Students can convert textbook images of small molecules into 3D representations in a molecular visualization program. (Amateur) AR2. Students can choose the best rendering of a macromolecule to use in a given situation. AR2.01 Students can recognize that a cartoon rendering is a summary of the detail in a line rendering. (Novice, Amateur) AR2.02 Students can describe the atoms and their representations in different renderings (e.g., coloring, showing hydrogens/double bonds). (Novice) AR2.03 Students can identify or create a suitable rendering, or combination of renderings, for a specific purpose (e.g., a surface rendering overlaid with a cartoon to highlight the van der Waals surface alongside secondary structure, or active site sticks shown over a cartoon). (Novice, Amateur) AR2.04 Students can identify the limitations in various renderings of molecular structures. (Amateur) AR2.05 Students can understand the level of detail of different molecular representations. (Novice, Amateur, Expert) AR2.06 Students can transition comfortably between equivalent 2D and 3D renderings of biomolecules. (Novice, Amateur, Expert) AR2.07 Students can use and interpret color in the context of macromolecules to clarify and/or highlight features (e.g., coloring amino acids differently by property, different molecules uniquely in a complex, protein chains, secondary structure). (Novice) Construction and Annotation (CA) Ability to build macromolecular models, either physical or computerized, and, where possible, add commentary, either written or verbal, to tell a molecular story. CA1. Students can compose information‐rich renderings of macromolecule‐ligand interactions. CA1.01 Students can construct and annotate a model of a macromolecule bound to a ligand. (Amateur) CA1.02 Students can construct a model of a macromolecule bound to a ligand and identify the types of molecular interactions. (Amateur) CA1.03 Students can construct a model of a macromolecule bound to a ligand and assess the importance of molecular interactions. (Expert) CA1.04 Students can produce a model of a macromolecule based on a known structure of a related macromolecule. (Amateur, Expert) CA2. Students can compose a rendering to predict the cellular location of a protein (e.g., extracellular, membrane associated, or cytoplasmic) based on the properties and orientations of functional groups. CA2.01 Students can design a rendering that conveys properties such as polarity, charge, secondary structure, etc. to suggest the cellular location of a macromolecule. (Amateur) CA2.02 Students can create protein images with colored polar/nonpolar residues to determine whether they fold with a hydrophobic core. (Amateur) CA2.03 Students can create images to display polar/nonpolar residues and propose a role for the protein and/or how it interacts with its environment ‐ and that the predictions would be plausible based on the protein. (Amateur) CA2.04 Students can make accurate predictions of the location/function of the protein that incorporates additional protein features, such as transmembrane helices, apparent docking surfaces, etc. (Expert) Ligands and Modifications (LM) Metals and metal clusters, additions such as glycosylation, phosphorylation, lipid attachment, methylation etc. LM1. Students can identify ligands and modified building blocks (e.g., hydroxyproline, aminosaccharides, modified nucleobase) within a rendered structure. LM1.01 Students can use the annotation associated with a pdb file to identify and locate ligands and modified building blocks in a given biomolecule. (Amateur) LM1.02 Students can visually identify non‐protein chemical components in a given rendered structure. (Amateur) LM1.03 Students can distinguish between nucleic acid and ligands (e.g., metal ions) in a given nucleic acid superstructure. (Amateur) LM1.04 Students can explain how a ligand in a given rendered structure associates with the biomolecule (e.g., covalent interaction with residue X). (Amateur) LM1.05 Students can locate/identify ligands and modified building blocks in unannotated structures and describe their role. (Expert) LM2. Students can describe the impact of a ligand or modified building block on the structure/function of a macromolecule. LM2.01 Students can look at a given rendered structure and describe how the presence of a specific ligand or modified building block alters the structure of that biomolecule. (Amateur) LM2.02 Students can explain how the removal of a particular ligand or modified building block would alter the structure of a given biomolecule. (Expert) LM2.03 Students can use molecular visualization tools to predict how a specified ligand or modified building block contributes to the function of a given protein. (Amateur, Expert) LM2.04 Students can predict how a ligand or modified building block contributes to the function of a protein for which the structure has been newly solved. (Expert) Macromolecular Assemblies (MA) Polypeptides, oligosaccharides, and nucleic acid and lipid superstructures (e.g. protein–nucleic acid complexes, lipid membrane-associated proteins) MA1. Students can describe various macromolecular assemblies. MA1.01 Students can identify individual biomolecules in a macromolecular assembly. (Novice, Amateur, Expert) MA1.02 Students can describe functions of individual biomolecules within a macromolecular assembly. (Novice, Amateur, Expert) MA1.03 Students recognize the various lipid ultrastructures (e.g., micelles, bicelles, vesicles, and lipid bilayers) in a 3D structure. (Novice) MA2. Students can compose information‐rich renderings of macromolecular assemblies. MA2.01 Students can render a macromolecular assembly to highlight individual structures. (Amateur) MA2.02 Students can render a macromolecular assembly to illustrate structural features (e.g., binding interfaces, symmetry, tertiary structure, etc.). (Novice, Amateur, Expert) Macromolecular Building Blocks (MB) Recognition of native amino acids, nucleotides, sugars, and other biomonomer units/building blocks. Understanding of their physical and chemical properties, particularly regarding functional groups. MB1. Students can identify individual building blocks of biological polymers. MB1.01 Given a rendered structure of a biological polymer, students can identify the ends of a biological polymer. (Novice, Amateur, Expert) MB1.02 Given a rendered structure, students can divide the polymer into its individual building blocks. (Novice) MB1.03 Given a rendered structure, students can identify the individual building blocks. (Novice) MB2. Students can describe the contributions different individual building blocks make in determining the 3‐D shape of the polymer. MB2.01 Students can describe the physical/chemical properties of an individual building block/functional group in a rendered structure of a polymer. (Amateur) MB2.02 Students can describe the significance of the location of individual building blocks within the 3D structure of a polymer (protein, carbohydrate, or nucleic acid). (Novice, Amateur, Expert) MB2.03 Students can identify physical/chemical properties of individual building blocks/functional groups in different local environments. (Amateur) MB2.04 Using a visualized structure, students can identify stereochemistry (e.g., in carbohydrate, lipid, and protein structures). (Amateur) MB2.05 Students can modify/mutate a building block to change the 3D structure of a polymer (protein, carbohydrate, or nucleic acid). (Amateur, Expert) Molecular Dynamics (MD) Animated motion simulating conformational changes involved in ligand binding or catalysis, or other molecular motion/dynamics. MD1. Students can describe the impact of the dynamic motion of a biomolecule on its function. MD1.01 Students can recognize that biological molecules have different conformations. (Novice, Amateur) MD1.02 Students can correlate molecular movement with function. (Novice, Amateur, Expert) MD2. Students can predict limits to macromolecular movement. MD2.01 Students can locate potential regions of flexibility and inflexibility in the structure of a biomolecule. (Novice, Amateur) MD2.02 Students can recognize acceptable/unacceptable movement within a macromolecule by determining whether the movement is within allowable bond angles. (Expert) MD2.03 Students can recognize acceptable/unacceptable movement within a macromolecule by determining whether the movement results in steric hindrance. (Amateur) MD2.04 Students can recognize acceptable/unacceptable movement within a macromolecule by considering the atomic packing constraints. (Expert) Molecular Interactions (MI) Covalent and noncovalent bonding governing ligand binding and subunit‐subunit interactions. MI1. Students can predict the existence of an interaction using structural and environmental information (e.g. bond lengths, charges, pH, dielectric constant). MI1.01 Students can distinguish between covalent and noncovalent interactions. (Novice) MI1.02 Students can identify different noncovalent interactions (e.g., hydrogen bonds, ionic interactions, van der Waals contacts, induced dipole) given a 3D structure. (Amateur) MI1.03 Students can predict whether a functional group (region) would be a hydrogen bond donor or acceptor. (Amateur) MI1.04 Students can render the 3D structure of a biomolecule so as to demonstrate the ionic interactions and/or charge distribution of the different non‐covalent interactions. (Amateur) MI1.05 As it relates to a particular rendered structure, students can rank the relative strengths of covalent and noncovalent interactions. (Amateur) MI2. Students can evaluate the effect of the local environment on various molecular interactions. MI2.01 Students can identify regions of a biomolecule that are exposed to or shielded from solvent. (Novice) MI2.02 Students can identify other molecules in the local environment (e.g., solvent, salt ions, metals, detergents, other small molecules) that impact a molecular interaction of interest. (Novice) MI2.03 Students can predict the impact of other molecules in the local environment (e.g., solvent, salt ions, metals, detergents, other small molecules) on a molecular interaction of interest. (Amateur) MI2.04 Students can predict the pKa of an ionizable group based on the influence of its local three-dimensional environment. (Amateur) MI2.05 Students can propose a change to the local environment that would yield a desired change in a molecular interaction. (Expert) MI2.06 Using molecular visualization tools, students can determine which intermolecular force is most critical to stabilizing a given interaction. (Expert) Symmetry/ Asymmetry Recognition (SA) Recognition of symmetry elements within both single chain and multi-chain macromolecules. SA1. Students can identify symmetric or asymmetric features in rendered molecules. SA1.01 Students can identify symmetric features in a rendered molecule (shown in fixed orientation). (Novice) SA1.02 Students can rotate a single macromolecule, multi-chain macromolecules (e.g., homo- or heteromers), complexes of macromolecules, and supramolecular assemblies to identify axes of symmetry. (Amateur) SA1.03 Students can identify symmetric and asymmetric features in rendered molecules after coloring a given rendered molecule to reveal structural features (charge, hydrophobicity, etc.). (Amateur) SA2. Students can hypothesize the functional significance of symmetry or asymmetry in rendered molecules. SA2.01 Students can explain the functional significance of rotational axes of symmetry (or asymmetry) in a given rendered molecule. (Novice, Amateur, Expert) SA2.02 Students can predict functional significance of symmetry (or asymmetry) in a given rendered molecule. (Amateur, Expert) Structure‐Function Relationship (SF) Active/binding sites, microenvironments, nucleophiles, redox centers, etc. (please also see LM2.03) SF1. Students can evaluate biomolecular interaction sites using molecular visualization tools. SF1.01 Students can identify functionally relevant cofactors, ligands or substrates associated with a macromolecule and describe their role (e.g., an active site magnesium ion). (Amateur, Expert) SF1.02 Students recognize that the size and shape of the ligand must match the size and shape of the binding site. (Novice, Amateur) SF1.03 Students recognize that the polarity or electrostatic potential of a surface complements that of the ligand or substrate. (Novice, Amateur) SF1.04 Students recognize that the hydrophobicity of a surface complements that of the ligand or substrate. (Novice, Amateur) SF1.05 REMOVED (integrated with SF1.03) SF1.06 Students can use docking software to predict how the surface properties of a macromolecule guide and allow the binding of a ligand or substrate. (Amateur) SF2. Using molecular visualization, students can predict the function of biomolecules. SF2.01 Students can recognize structurally related molecules. (Novice) SF2.02 Students can superimpose structurally related molecules. (Novice, Amateur) SF2.03 Students can identify functionally relevant features of a macromolecule (e.g., an active site cysteine, a functional loop). (Amateur) SF2.04 Students can predict molecular function given a binding site. (Amateur, Expert) SF3. Using molecular visualization, students can predict the function of an altered macromolecule. SF3.01 Students can structurally alter a macromolecule. (Novice) SF3.02 Students can propose structural alterations to test interactions in a macromolecule. (Amateur) SF3.03 Students can predict the impact of a structural alteration on the function of a macromolecule. (Amateur, Expert) Structural Model Skepticism (SK) Recognition of the limitations of models to describe the structure of macromolecules. SK1. Students can critique the limitations of a structural model of a macromolecule. SK1.01 Students can explain that the pdb file is a model based on data and that, as a model, it has limitations. (Novice, Amateur) SK1.02 Students associate resolution with reliability of atom positions. (Amateur) SK1.03 Students can identify building blocks (for example, amino acid side chains) whose orientation in a biopolymer is uncertain. (Expert) SK1.04 Students can evaluate the flexibility/disorder of various regions of a macromolecular structure. (Novice, Amateur, Expert) SK1.05 Students can reconcile inconsistent numbering of individual building blocks among species and structure files. (Novice) SK1.06 Students can utilize a Ramachandran plot/steric clashes to interpret the validity of a structure. (Amateur, Expert) SK1.07 Students can describe the limitations of a macromolecule‐ligand docking simulation. (Amateur, Expert) SK2. Students can evaluate the quality of 3D models including features that are open to alternate interpretations based on molecular visualization and PDB flat files. SK2.01 Students can evaluate a crystal structure for crystal packing effects. (Novice, Amateur, Expert) SK2.02 Students can resolve differences between the asymmetric unit and the functional biological assembly. (Expert) SK2.03 Students can differentiate functional ligands (with biological/biochemistry role) from nonfunctional ligands (most solvents, salts, ions, and crystallization agents). (Novice, Amateur, Expert) SK3. Students can discuss the value of experimentally altering a biomolecule to facilitate structure determination. SK3.01 Students can identify non‐native structural features. (Amateur) SK3.02 Students can propose molecular modifications to facilitate structure determination. (Amateur, Expert) SK3.03 Students can propose a purpose for the introduction of non‐native structural features to facilitate structure determination. (Amateur, Expert) Topology and Connectivity (TC) Following the chain direction through the molecule, translating between 2D topology mapping and 3D rendering. TC1. Students can describe or illustrate the linkages between building blocks within a macromolecule. TC1.01 Students can trace the backbone of a macromolecule in three dimensions. (Novice, Amateur) TC1.02 Students can use appropriate terms to describe the linkages/bonds/interactions that join individual building blocks together in a macromolecule or macromolecular assembly. (Novice, Amateur) TC1.03 Given a virtual model of individual building blocks, students can predict the types of linkages/bonds/interactions that are possible or favorable. (Amateur) TC1.04 Given individual building blocks, students can appropriately connect them to create a biological polymer (e.g., drawing carbohydrate linkages, a small peptide). (Amateur, Expert) TC2. Students can describe the overall shape and common motifs within a 3D macromolecular structure. TC2.01 Using molecular visualization software, students can describe the three-dimensional structure of a macromolecule, including overall shape and common structural motifs. (Novice, Amateur, Expert) TC2.02 Students can identify common domains/motifs within a macromolecule. (Amateur, Expert) TC2.03 Students can identify connectivity features between domains or subunits in a macromolecular structure. (Amateur) TC2.04 Students can identify interactions between domains or subunits in a macromolecular structure. (Amateur, Expert) TC2.05 Students can describe how domains/motifs in a macromolecule work together to achieve a concerted function in the cell. (Amateur, Expert) TC2.06 Students can identify the levels of protein structure (e.g., parse a tertiary/quaternary structure into a series of secondary structures/motifs) and the ways in which they are connected from a three‐dimensional structure. (Novice, Amateur, Expert) TC3. Students can explain how any given biomolecular interaction site can be made by a variety of topologies. TC3.01 Students can recognize that the groups that comprise a functional site only require proper arrangement in three-dimensional space rather than a particular order or position in the linear sequence. (Amateur) TC3.02 Students can recognize similarities and differences in two similar ‐ but not identical ‐ three dimensional structures. (Amateur) TC3.03 Students can describe dissimilar portions of homologous proteins as arising from genetic insertions/deletions/rearrangements. (Amateur)

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/iCn3D_for_Biomolecular_Visualization_Learning_Themes_and_Goals/BioMolViz_Framework.txt

It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, BioMolViz Theme: Atomic Geometry (AG) It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, BioMolViz Theme: Construction and Annotation % It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, BioMolViz Theme: Ligands and Modifications ( It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, BioMolViz Theme: Macromolecular Assemblies ( It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, BioMolViz Theme: Macromolecular Building Block It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, BioMolViz Theme: Molecular Dynamics (MD) It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, BioMolViz Theme: Molecular Interactions (MI% It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, BioMolViz Theme: Structural Model Skepticism % It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, BioMolViz Theme: Structure‐Function Re It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, Asymmetry Recognit It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, BioMolViz Theme: Topology and Connectivity ( It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz, Biomolviz- Constructing iCn3D Models to Target B It is not only important to visualize pre-rendered models of biomolecules, but it is also important to be able to create them to address key aspects of structure and function.  These efforts should be guided by a clear set of learning goals and objectives that target student understanding of structure and function.  So it is fortunate that clear learning themes, goals and objectives are articulated in a Biomolecular Visualization Framework created by BioMolViz,

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Fundamentals_of_Biochemistry_Vol._V_-_Problems/iCn3D_for_Biomolecular_Visualization_Learning_Themes_and_Goals/BioMolViz_Theme%3A_Alternate_Renderings_%28AR%29.txt

Search Fundamentals of Biochemistry Introduction to Cell Signaling Cell signaling is at the heart of biology. A cell must know how to respond to chemical signals in its environment. These signals control every aspect of cell life and interactions. A cell must sense when to grow, divide and die. It must sense the presence of foreign and toxic molecules. It must defend itself. The membrane represents the divide between the outside and inside world. Signals must cross that divide and this most often happens without the signaling molecule entering the cell. Just the signal itself is transduced across the membrane. And it doesn't stop there. Internal signaling, also across internal membranes of organelles in eukaryotes, propagates spatially and temporally across the cytoplasm in all cells (prokaryotes, Archaea, and eukaryotes). It's impossible to describe the myriad of processes that occur in cell signaling in a single chapter, but we will do our best to present the common features used across cells. In addition, signals in complex organisms must be integrated within tissues and organs, between organs (brain and liver for example), and within the entire organisms. Mathematical modeling is critical in understanding the complexity of these interconnected interactions. Consider the flee, fight, or freeze responses. What if you were walking down the street and suddenly saw a tiger approaching you? Most would flee. In that process, the neural, muscular, and metabolic systems must be integrated. The resting state is followed by the activation of the flight/flee response, the maintenance of the fleeing state, and then the return to the resting state. An alternative is the freeze response, which in some situations would be adaptive. Perhaps the most complex signaling occurs in the great communication networks in the neural and immune systems. Think of it. You can experience a traumatic or emotionally-charged event once and remember it forever. Ordinary events leave less permanent traces. The biochemical changes accompanying long and short-term memory are fascinating Let's consider general principles for signal transduction across membranes of any cell that must respond to its environment. Typically the agent that signals a cell to respond is a molecular signal. Signaling can also be mediated by pressure (for touch and hearing) or light for vision. The chemical signal binds either to a cell surface receptor or to a cytoplasmic receptor if the signaling agent is hydrophobic and can transit the membrane bilayer. The logic of signaling To a first approximation, you could consider a cell as a black box, as shown in Figure \(1\), with input (an external molecule for example) and resulting output signals within a cell, such as activation of gene transcription, trafficking of molecules within the cell, chemical reactions, or even export of molecules from the cell. There could be one or more signals, and one or more outputs. The inputs might arrive gradually and reach a threshold before triggering an output, or the input might be abrupt leading to an immediate output. This Figure \(1\): Note the use of terminology taken from the world of electronics. Figure \(2\) shows the similarity in the complexity of an electronic wiring diagram (left) to the interconnected metabolic pathways of a cell (right). The analogy is much greater than you might think. Cell signaling researchers have adopted the language and symbols of electronics as they consider how two inputs could lead to different output signals. Figure \(3\) shows a truth table with different Boolean operator logic gates and symbols used by electrical engineers that also apply to signals to cells and their outputs. Two input signals, A and B, arrive at different logic gates named AND, NOR, OR, XOR AND, and NAND. 0 indicates no signal (either input or output) while 1 indicates a signal (either input or output). Each of these logic gates leads to different outputs: • AND gates require both A and B input signals for an output; • OR gates require either A or B or both for an output signal; • NOR gates require neither A or B for an output signal; • NAND gates normally have an output signal unless both inputs A and B are present; • XOR (Exclusionary OR) gates give a signal if the two inputs A and B differ. An example of an analogous biological AND gate is N-WASP protein [a homolog to the Wiskott-Aldrich syndrome protein (WASP)]. This protein regulates actin polymerization and binds multiple signals. Stimulation with two proteins, Cdc42 and phosphatidylinositol (4,5)-bisphosphate (PIP2), activates polymerization. Researchers are designing protein logic gates by creating a series of heterodimeric proteins. Consider the following heterodimers: A:A', B:B', and C:C', where the second member in each pair is different from the first and bound reversibly through noncovalent interactions (indicated by the colon :) Other versions could exist, such as A:C'. An AND gate for the formation of an A:C' dimer would be made by making the covalent, single molecules A'-B and B'-C. The A:C' dimer could form only in the presence of both A'-B and B'-C. This is illustrated in Figure \(4\). Signals and their responses must occur at the right time under the right conditions and for the right duration. Under opposing sets of conditions (for example well feed and starving), opposing signaling pathways, mediated by different signaling molecules, must be mutually integrated and regulated so one is turned on and one shut off. Methods must be in place to terminate signal effects. Given the complexity of signaling pathways, it's hard to know how to present the material in a single chapter. Binding initiates and mediates almost all biological events. That binding must also be specific to avoid off-target effects. If you were to predict what biomolecule could confer specificity in the binding of a signal and allow changes from the unbound to bound state, you would certainly pick proteins. Protein:ligand binding is key to understanding biosignaling. Other types of biomolecules such as lipids and nucleic acid are involved and are of clear importance, but they are typically involved downstream from the initial protein:signal binding event. Hence we will focus mostly on signaling proteins and conformational/activity changes that occur on signal binding. To simplify protein involvement in signaling, we can assume signaling proteins have an active and inactive state. The states are interconvertible (reversible). Let's assume the signaling protein is an enzyme. The activity of the enzyme depends on many factors as described in Figure (5\) below. The green color represents the active enzyme, while red indicated inactive. Of course, proteins that are not enzymes (for example transcription factors) can be regulated similarly. In addition, the amount (and localization as well) of a signaling protein can regulate the activity of the protein, as illustrated in Figure (6\) below. Each signaling protein can be considered to be a node in a larger pathway consisting of interconnected nodes. This chapter on cell signaling hence can be viewed as a capstone to Unit 1, which explores the structure and function of biomolecules and as a prelude to the study of whole metabolic pathways. Second Messengers and Signal Translocation The chemical species that trigger signaling typically binds to a target protein transmembrane receptor on the surface of a cell but does not itself enter the cell. Just the signal enters the cell. If you were to hypothesize how that might happen, you would predict two mechanisms: • an integral transmembrane receptor undergoes a conformational change on binding that propagates to its cytoplasmic domain, which can then interact either with other cytoplasmic proteins or cytoplasmic domains of other membrane proteins, transmitting the symbol into the cytoplasm; • the bound conformation of the receptor has enzyme activity and can catalyze chemical reactions on the luminal side of the membrane. This could include the chemical modification of proteins or the synthesis of new small molecules from cell metabolites. These new small molecules are called second messengers. They could be formed in the membrane bilayer or the cytoplasm. There are several diverse types of second messengers. Two common ones are cyclic derivatives of small nucleotides, including cyclic AMP (cAMP) derived from ATP and cyclic GMP (cGMP) derived from GTP. Ca2+ ions are usually found in low concentrations in the cytoplasm as they are pumped into internal organelles such as the endoplasmic reticulum and mitochondrion. Signaling processes can release Ca2+ ions in waves within the cell. Membrane lipids are also processed to form second messengers. Membrane phospholipids can be cleaved by cell signaling-activated lipases to form free arachidonic acid, sphingosine, diacylglycerol, and inositol-trisphosphate, which can act as second messengers. Redox signaling in the cell can also occur through hydroperoxides acting as second messengers. What if the response of the cell requires gene transcription? Somehow the signal has to translocate from the cell membrane through the cytoplasm through the nuclear membrane into the nucleus. Hence a series of translocations of multiple downstream signaling events must occur. We have already seen how newly synthesized proteins have signal sequences that target them to specific cellular locations like the cell membrane, mitochondria, nucleus (nuclear localization sequences - NLS and the small GTP binding protein RAN), or for export. Proteins involved in signaling also can move throughout the cell as part of the signaling process. Likewise, we have seen how cytoplasmic proteins can be targeted to membranes by attachment of fatty acids or isoprenoids. Post-translational modification of signaling proteins Nature has chosen the post-translational modifications (PTM) of proteins as a ubiquitous way to alter the signaling states of proteins. As we have seen previously, PTMs can alter protein conformation. They can also present new binding interfaces that allow interaction with other signaling proteins. The main (but not the only) PTM used for signaling is the reversible phosphorylation of the OH-containing amino acid side chains (Tyr, Ser, and Thr) as well as histidine (mostly in prokaryotes), so we will focus on them. Enzymes that catalyse the phosphorylation of proteins are called protein kinases. Reversibility is important since if phosphorylation of a target protein is associated with a specific signaling change (either activation or inhibition), then dephosphorylation can easily reverse the signaling event. Enzymes that dephosphorylate phosphoproteins are called protein phosphatases. Figure \(7\) shows the generic reaction of protein kinases and phosphatases. There appear to be 518 protein kinases and 199 protein phosphatases encoded in the human genome. Why so many? If just one protein kinase existed, one mutation in it would be disastrous. In addition, the large number of kinases and phosphatase allows for great control in the specificity of these enzymes for their target protein substrates. Kinases Kinases are a class of enzymes that use ATP to phosphorylate molecules within the cell. The names given to kinases show the substrate which is phosphorylated by the enzyme. For example: • hexokinase - an enzyme that uses ATP to phosphorylate hexoses. • protein kinase - enzymes that use ATP to phosphorylate proteins within the cell. (Note: Hexokinase is a protein, but is not a protein kinase). • phosphorylase kinase: an enzyme that uses ATP to phosphorylate the protein phosphorylase within the cell If a protein is phosphorylated by a kinase, the phosphate group must eventually be removed by a phosphatase through hydrolysis. If it wasn't, the phosphorylated protein would be in a constant state of either being activated or inhibited. Kinases and phosphatases regulate all aspects of cellular function. About 1-2% of the entire genome encodes kinases and phosphatases. Kinases can be classified in many ways. One is substrate specificity: Eukaryotes have different kinases that phosphorylate serine/threonine or tyrosine side chains. Prokaryotes also have His and Asp kinases, but these are unrelated structurally to the eukaryotic kinases. There are 11 structurally different families of eukaryotic kinases, which all fold to a similar active site with an activation loop and catalytic loop between which substrates (ATP and the OH-containing side chain) bind. Simple, single-cell eukaryotic cells (like yeast) have predominantly cytoplasmic Ser/Thr kinases, while more complex eukaryotic cells (like human cells) have many Tyr kinases. These include the membrane-receptor Tyr kinases and the cytoplasmic Src kinases. Manning et al. have analyzed the entire human genome (DNA and transcripts) and have identified 518 different protein kinases, which cluster into 7 main families as shown in Table \(1\): below. Family membership was determined by sequence comparisons of catalytic domains. The entire repertoire of kinases in the genome is called the kinome. Alterations in 218 of these appear to be associated with human diseases. Name Description AGC Contain PKA, PKG, and PKC families CAMK Ca2+/CAM-dependent PK CKI Casein kinase 1 CMGC Contain CDK, MAPK,GSK3, CLK families STE homologs of yeast sterile 7, 11, 20 kinases; MAP Kinase PTK Protein tyrosine kinase PTKL Protein tyrosine kinase-like RGC Receptor guanylate kinase Table \(1\): The human kinome Phosphatases There are three main families of phosphatases, the phospho-Tyr phosphatases (PTP), the phospho-Ser/Thr phosphatases, and those that cleave both. Of all phosphorylation sites, most (86%) are on Ser, 12% involve Thr, and about 2% on Tyr. They can also be categorized by their molecular sizes, inhibitors, divalent cation requirements, etc. In contrast to kinases which differ in the structure of their catalytic domains, many phosphatases (PPs below) gain specificity by binding protein cofactors which facilitate translocation and binding to specific phosphoproteins. The active phosphatase hence often consists of a complex of the phosphatase catalytic subunit and a regulatory subunit. Regulatory subunits for Tyr phosphatases may contain a SH2 domain allowing binding of the binary complex to autophosphorylated membrane receptor Tyr kinases. Given this background, we can now start to explore signal transduction in a methodical way. There are two major ways to organize our discussions: • start from the binding of the molecular signal at the cell membrane and trace the signaling events inward into the cell, potentially all the way to the nucleus and gene expression • describe recurring motifs found in most pathways. We will use a combination of both, but it makes sense to start with the signaling proteins at the cell membrane. Next, we will focus on second messengers. We'll follow that with detailed explorations of kinases and phosphatases and specific signaling pathways.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Unit_IV_-_Special_Topics/28%3A_Biosignaling_-_Capstone_Volume_I/28.01%3A_General_Features_of_Signal_Transduction.txt

Search Fundamentals of Biochemistry Introduction We are going to start a more detailed description of cell signaling where it begins, at the cell membrane, and move inward, toward intracellular organs, where we will end at the nucleus and changes in gene expression mediated by the signal. Of course, this end is somewhat arbitrary as the signal could propagate from the nucleus back to the membrane where newly synthesized membrane protein might be inserted or even exported, as in the case of secreted antibodies. It is difficult enough to keep track of all the molecular players, let alone add onto that their initial location in the cell and their final location if they translocate. Signaling can also be more daunting for those with a more chemistry focus, who find the details of cellular structure and trafficking a bit daunting. Figure \(1\) shows a truncated view of the cell membrane, membrane proteins, and some of the organelles that we will visit throughout this chapter. To stay localized we will repeat the figure or variants of it several times in this section. Receptors at Cell Membrane Let's start at the location where signaling almost invariable begins, in the cell membrane, where signals (hormones, neurotransmitters, nutrients, other cells) bind to cell surface transmembrane proteins shown in the red box in Figure \(2\). We have already discussed integral and peripheral membrane proteins in Chapter 11. When occupied by a ligand signal, the signaling event mediated by a transmembrane receptor might be a change in the membrane protein conformation, which propagates to its intracellular domain. Alternatively, the receptor might change its conformation and become a ligand-gated kinase (or possibly a phosphatase) or a ligand-gated channel or pore. We will discuss signal-gated ion channels in Chapter 28.9 - Neural Signaling. Ultimately, intracellular enzymes are activated in cells in response to the external molecular signal. This provides amplification of the initial signal since a single activated enzyme undergoes multiple rounds of catalysis before it would become inactivated. If the multiple products (a second messenger or phosphorylated proteins for example) that are formed activate multiple addition enzymes, the signal is further amplified. Receptors with no kinase or transport activity - G Protein-Coupled Receptors (GPCRs) One major type of signaling receptor is the G-protein coupled receptor (GPCR). Over 800 GPCRs are encoded in the human genome which represents almost 4-5% of the number of protein-coding genes. The proteins below to five major families: rhodopsin, secretin, glutamate, adhesion, and frizzled/taste2. They don’t express enzymatic activity but on binding, they can activate enzymes inside the cell by interacting through their cytoplasmic domains with G proteins (GTP binding proteins) in the cytoplasm. GPCRs have been called serpentine receptors as their single amino acid chain proteins have 7 transmembrane-spanning α- helices. All of the GPCRs have similar yet slightly different structures, allowing them to interact with specific ligands. Many of the GPCRs bind to unknown ligands and hence are called orphan receptors. We will explore a few GPCRs in more detail below. GPCRs that modulate the membrane enzyme adenylyl cyclase: β-adrenergic receptor The β-adrenergic receptor is a prototypical GPCR. Found in muscle, liver, and fat cells, it binds epinephrine and adrenaline, which leads to energy mobilization and muscle activation (i.e. flight or fight response). The mechanism of activation of a GPCR is illustrated using the beta-adrenergic receptor as an example. The unoccupied adrenergic receptor is associated with a heterotrimeric G protein, which contains an α, β, and γ subunits. GDP is usually bound to the α subunit. Figure \(3\) shows a cartoon of the GPCR interacting with the heterotrimeric G protein. When the hormone is bound to the receptor, a conformation change is propagated to the cytoplasmic domain of the GPCR altering its interaction with Gαβγ. This causes a conformational change in the Gα subunit, which leads to an exchange of bound GDP with GTP in the Gα subunit, promoting the dissociation of the Gα-GTP from Gβγ. Gα-GTP then binds to and activates an adjacent membrane enzyme called adenylate cyclase, which produces a second messenger by converting ATP to cyclic AMP (cAMP). Figure \(4\) shows steps in the generic GPCR activation cycle. Figure \(4\): Activation cycle of G-proteins by G-protein-coupled receptors. https://commons.wikimedia.org/wiki/F...PCR-Zyklus.png Creative Commons Attribution-Share Alike 3.0 Unported The primary message binds to the GPCR (1 leading to state 2. The cytoplasmic domain conformational changes allow GTP exchange in the Gα subunit as state 3 goes to 4. In 5 the Gα-GTP complex dissociates from Gβγ, which remains in the membrane. The Gα-GTP subunit is held and localized to the membrane (not evident in the above figure) through a lipid anchor attached through a post-translational modification. Remember, the GPCR has no ligand-gated enzymatic activity. Yet it indirectly leads to the activation of a membrane enzyme, adenylate cyclase, when the dissociated Gα-GTP binds to the cyclase. As long as GTP remains bound to the Gα subunit, it will continue to modulate the activity of adenylate cyclase. A built-in regulatory mechanism does exist in the protein since the Gα subunit has GTPase activity. The GTP will eventually hydrolyze, and the GDP-Gα subunit will lose affinity for its bound partner (adenylate cyclase), and return to the heterotrimeric G protein associated with the unbound receptor. GPCRs bind the signaling ligand (primary message) in a binding cavity localized at the extracellular face and between four of the transmembrane helices. The activity and structure of GPCRs have been studied using natural ligands (hormones and neurotransmitters), as well as agonists, partial agonists, inverse agonists, and antagonists. As discussed previously, agonists bind to the natural ligand binding site and elicit the same or a partial response (partial agonist). Inverse agonists bind and lower the response of a constitutively active receptor, and antagonists bind and prevent the normal response of an agonist. About 35% of pharmaceutical drugs target GPCRs, but target only about 15% of the ∼800 human GPCRs. The orphan GPCRs are increasingly targets for drug development. Most hormones and neurotransmitters work through GPCRs. In addition, our primary senses of vision, smell (olfaction), and taste (gustation) work through GPCRs. Figure \(5\) shows the structure of the beta 2-andrenergic:Gs complex with bound agonist. No membrane is shown. The GPCR is shown in cyan. The 7 transmembrane helices should be obvious. The ligand is bound between them. The Gα subunit is shown in magenta, the Gβ in dark blue, and the Gγ in orange/brown. The gray subunit is a Camelid antibody VHH fragment (a single-chain nanoantibody used to stabilize the conformation for crystallization. ) The biggest conformational change that occurs when the GPCR binds an agonist is an outward movement (14 Å) at the intracellular domain of transmembrane segment 6 (TM6) and an extension of the TM5 helix. This leads to a movement of the Gα's alpha-helical domain, enabling GTP exchange for GTP. Of course, multiple reactions determine the fraction of the Gα in the active GTP-bound state. These would include the relative cellular concentrations of free cytoplasmic GDP and GTP, their KD values for the Gα, the rate constant for the hydrolysis of bound GTP, and the rate constants for the GDP ↔ GTP exchange. Figure \(6\) shows an interactive iCn3D model of the Crystal structure of the beta2 adrenergic receptor-Gs protein complex 3SN6 The GPCR is shown in green, the Gα in magenta, Gβ in blue, and the Gγ in brown. An agonist (spacefill) is shown in CPK colors near the outer leaflet. Some bacterial toxins work by inactivating the GTPase activity of the Gα subunit, keeping it in the "stuck" position. For example, cholera toxin, an enzyme released by Vibrio cholera, catalyzes the ADP ribosylation of an Arg in the Gα subunit by transferring everything but the nicotinamide from NAD+ to the Arg residue. Since the Gα subunit stimulated the activity of adenylyl cyclases, it is often named a stimulatory Gα protein or G. Figure \(7\) shows an interactive iCn3D model of the adenylyl cyclase activator G with GTP-γ-S (1azt) Now we can explore how the occupied GPCR, which again has no enzymatic activity, activates the enzyme adenylyl cyclase. Figure \(8\) shows a cartoon of the G subunit bound to adenylyl cyclase as the Gβγ heterodimer remains associated with the membrane Adenylate cyclase converts ATP into the second messenger cyclic AMP (cAMP) as shown in Figure \(9\). The figure also shows how cAMP is broken down into AMP by the enzyme cAMP-specific 3',5'-cyclic phosphodiesterase (PDE). The latter step is necessary to control the lifetime of the second messenger cAMP. Why cAMP? You might ask why cAMP and not just AMP is nature's choice for GPCR signaling. AMP is a very important metabolic species. High concentrations of it signal an energy-depleted state. AMP is used in another signaling process to mobilize a response to adjust the energy state of a cell using a protein called AMP-Protein Kinase, which we will describe later. Many enzymes are also allosterically regulated by AMP. Figure \(10\) shows an interactive iCn3D model of the membrane adenylyl cyclase bound to an activated stimulatory G protein 6R3Q determined by cryo-EM. The carboxyl-terminal cytoplasmic domain has the catalytic and allosteric sites. Cannabinoid Receptors In contrast to the beta-adrenergic receptor which mediates activation of adenylate cyclase through G, some Gα subunits inhibit adenylate cyclase when bound. These Gα subunits are called Gi/oα in contrast to the stimulatory subunits, G. Also, Gα subunits interact with many proteins other than adenylate cyclase. Examples include cannabinoid receptors. Cannabinoid receptors are named after the exogenous and psychoactive drug Δ9-tetrahydrocannabinol (THC) that binds to the receptor. THC is the major phytocannabinoid (from plants) found in the Cannabis sativa plant and the marijuana-derived from it. The other main cannabinoid in the plant is cannabidiol (CBD). Phytocannabinoids bind to two types of human cannabinoid (CB) receptors, CB1 and CB2. They have 44% amino acid and 68% homologies in the entire protein and the transmembrane domain, respectively. The phytocannabinoids exert their effect through binding to human CB1 and CB2, whose endogenous ligands are two fatty acids derivatives called anandamide (AEA) and 2-arachidonoyl glycerol (2-AG). The structure of THC, CBD, and the two major endogenous ligands are shown in Figure \(11\). Figure \(11\): Structure of agonist and antagonist for cannabinoid receptors The cannabinoids from Cannabis sativa have a monoterpene isoprenyl group (C10) and a pentyl side chain (C5). The ligands are largely hydrophobic and probably access their binding site in the receptor mainly by lateral movement in the membrane. The receptors differ most in the N-terminal extracellular loop which is also involved in ligand binding. Figure \(12\) shows an interactive iCn3D model of the class A GPCR Cannabinoid Receptor-Gi Complex Structure with bound agonist (6KPF) The gray protein is a nanobody used to stabilize the protein during crystallization. The agonist is AM12033, which is similar to AM11542 in the figure above, but with a -C=N terminus instead of a bromide. THC and CBD Cannabis sativa contains the psychoactive drug, Δ9-tetrahydrocannabinol (THC), which is a partial agonist for CB1 (binds with reported Ki values of 10 or 53 nm)and CB2 (Ki = 40 nm). Its psychoactive effects on mental activity as well as pain and appetite are well known. In contrast, cannabidiol (CBD) is the main, non-psychoactive cannabinoid. It has a much lower affinity for the recombinant CB1 (Ki = 1.5 µM ) and CB2 (Ki = 3.7 µM). It appears to be a partial antagonist for CB1 and a weak inverse agonist for CB2. It has also been shown that CBD is a negative allosteric modulator of the agonistic effects of THC and 2AG. The actual psychotropic effects of combining THC and CBD are complicated and not understood well. Figure \(13\) shows an interactive iCn3D model of human CB1 in complex with agonist AM11542 (5XRA) How much of a receptor is bound with a cannabinoid depends on the concentration of the cannabinoid ligand and the Ki for the drug. The amount of THC and CBD depends on the genetics of the plant, which has been engineered to greatly decrease THC production (in the hemp plant used for nonpharmacological commercial properties) or increase either THC or CBD production at the expense of the other. The synthesis of THC and CBD proceeds through a common precursor, CBGA (cannabigerol acid). Two key flavoproteins, Δ9-tetrahydrocannabinolic acid synthase (THCAS) and cannabidiolic acid synthase (CBDAS) convert this common precursor CBGA into two new precursors, Δ9-THCA and CBDA, respectively. This final synthetic step involves an oxidative cyclization reaction using O2 and produces H2O2. Spontaneous, non-catalyzed decarboxylation and rearrangements of Δ9-THCA and CBDA lead to the final products, THC and CBD. This last process occurs on exposure to heat, which occurs in smoking and baking, and at a slower rate during storage. Commercially used preparations of THC and CBD for medicinal purposes vary widely in concentrations. THC concentration ranges for pain management (<5-10%) are much lower than those for psychotropic effects (<15%), with values of 21% or often found in "recreational" cannabis. High-potency THC strains can contain up to 25-30% THC by dry weight. For strands modified for CBD production, the maximal amount is about 25%. Even though CBD appears to be a partial antagonist for CB1, it appears that the ratio of THC:CBD is important in modulating the "high" or intoxicating state of THC. A ratio of THC:CBD of just over 1:1 leads to synergism or enhancement of the acute effects of THC whereas ratios of THC:CBD of 1:2 to 1:6 seem to have the least intoxicating effects. However, CBD decreases psychotic symptoms of THC and also decreases memory changes associated with THC. CBD also is an allosteric modulator of the μ-opioid receptor. At present, there are no structures of CBD bound to its cannabinoid receptors. Figure \(14\) shows an interactive iCn3D model of the CBD-bound full-length rat transient receptor potential vanilloid 2 (TRPV2) in nanodiscs (6U88) Now let's make it even more complicated. There are more than 20 different Gα-like proteins known in 4 major families. • Gs and Gi regulate adenylyl cyclase • Gq activates phospholipase Cβ (described below). There are 4 members given these strange names: Gq, G11, G14, and G15/16 • G12/13 activate small GTPase protein (described in Chapter 28.5) The Gα protein involved in light sensation is named transducin, while those involved in odorant detection and taste are called Golfactory and Ggustatory, respectively. As we add more variants of each signaling component, the origin of the complexity of signaling systems becomes evident. For example, the neurotransmitter serotonin binds to its receptor, a GPCR, and instead of gating the protein open to ion flow (as with other ligand-gated ion channels in the activation of neurons as we will see in Chapter 28.9), it interacts with two different alpha subunits, Gs, which leads to activation of adenylyl cyclase, and G12, which interacts with other small GTP binding proteins called GEFs (we will also see these later). GPCRs modulate the activity of the membrane enzyme phospholipase C. These receptors use the same mechanism for activation of the membrane enzyme adenylyl cyclase. When the primary signal is bound to the GPCR, a conformation change is propagated to the cytoplasmic domain of the GPCR, altering its interaction with Gαβγ in which the alpha subunit is a member of the Gα(q) family. This causes a conformational change in the Gα subunit which leads to an exchange of bound GDP with GTP in the Gα subunit, promoting the dissociation of the Gα-GTP. Gα-GTP then binds to and activates an adjacent membrane enzyme, phospholipase C (PLC), which cleaves membrane phospholipids to produce two, second messengers, diacylglycerol and inositol 1,4,5-trisphosphate (IP3). Their structures are shown in Figure \(15\). There appear to be 13 kinds of mammalian phospholipase Cs divided into six isotypes (β, γ, δ, ε, ζ, η). Phospholipase C is also named 1-phosphatidylinositol 4,5-bisphosphate phosphodiesterase. Figure \(16\) shows an interactive iCn3D model of the Gα(q)-phospholipase C-β 3 structure (4GNK). The magenta structure is the Gα(q) protein with bound GDP in spacefill. The cyan structure is the Pleckstrin Homology (PH) domain of the protein. This domain targets proteins to inositol phospholipids in the membrane but appears not to have this function in this protein. Note that in contrast to adenylyl cyclase, PLC is a peripheral, not an integral membrane protein. It is found in the cytoplasm as well as associated with the inner leaflet of the cell membrane where its main activities, regulating and cleaving PIP2, occur. PLC localizes to lipid rafts enriched in PIP2. Figure \(17\) shows the domain structure of phospholipase C β3 The N-terminal PH_14 represents the Pleckstrin Homology (PH) domain, which is among the top 15 of all domains in the human genome. All PLCs except PLCζ have this domain. Note that in another example of complexity, PLC-β binding to the inner leaflet does not require PIP2. The iCn3D model above indeed shows binding to the membrane seems to depend on adjacent structures and not the PH domain (cyan) specifically. Tabel \(1\) below characteristics of common signals that signal through GPCRs. signal vasopressin epinephrine light odorant odorant sweet tastant receptor VR β-adrenergic rhodopsin odorant receptor 1 odorant receptor 2 sweet receptor Ga-like subunit Gi Gs transducin Golfactory Golfactory Ggustatory coupled enzyme adenylate cyclase adenylate cyclase phosphodiesterase phospholipase C adenylate cyclase adenylate cyclase 2nd messenger decrease cAMP increase cAMP decrease cGMP increase IP3 increase cAMP increase cAMP protein affected decrease PrK-A increase PrK-A dec. Ca, Na perm. inc. Ca perm inc.Ca, Na perm dec. K perm Tabel \(1\): Characteristics of common signals that work through GPCRs. Changeux and Edelstein reviewed the MHC model 40 years after its conception and support its application to signal transduction processes. They include in signaling molecules not only hemoglobin, but regulatory enzymes (aspartate transcarbamylase, phosphofructokinase, LDH, glycogen phosphorylase), membrane receptors (acetylcholine receptor, rhodopsin), and nuclear receptors (lac repressor, steroid hormone receptors). In all these signaling proteins, residue distant from the "active" site participates in binding to allosteric ligands. Often the allosteric site is on a separate domain that can be cleaved from the protein and still maintain allosteric ligand binding properties. The proteins also consist of multiple subunits easily related by distinct symmetry axes. Allosteric ligands often bind in cavities in subunit interfaces along symmetry axes. In general, crystal structure analyses show that low-affinity T and high-affinity R forms of the signaling proteins exist but are accompanied by minor tertiary structure changes in individual subunits (i.e. perfect symmetry in all subunits is not preserved on the binding of allosteric ligand). For neurotransmitter membrane receptors, these two states can be correlated with an open and closed state (for ion flux), and open conformations of these proteins can often be found in mutant forms. However, for many ligand-gated ion channels and G-protein coupled receptors (serpentine), kinetic analyses show more complicated forms than can be represented by a simple two-state (R and T) model. High-resolution microscopy shows evidence for nonsymmetrical quaternary structural changes. These changes can be observed in the absence of ligand, which gives support to the MWC concept that allosteric ligands select certain conformational states, leading to equilibrium shifts in the unliganded receptor to the more high-affinity state. More refined methods of structural analysis will presumably show more evidence of subtle tertiary changes in the proteins that are preludes to quaternary structural changes. Yet the simplicity of the MWC model for explaining many features of signaling proteins remains. Receptors with signal-gated kinase activity - Receptor Tyrosine Kinases (RTK) Why bother binding a primary message to a GPCR and going through multiple steps before the activation of a membrane enzyme like adenylate cyclase? Wouldn't it be easier and more efficient to have the membrane receptor a ligand-activated enzyme? Such is the case with special membrane receptors called Receptor Tyrosine Kinases (RTKs). There are about 90 tyrosine kinases in the human genome of which 58 are RTKs. Figure \(18\) shows the family domain structure of the RTKs. Note that the insulin receptor (InsR) is a dimer of two monomeric insulin receptor chains. Figure \(19\)s shows in more detail the domain structure of the epidermal growth factor receptor (EGFR). Figure \(19\): Domain structure of the EGFR domain. The domain (red/brown) immediately after the blue domain is the transmembrane domain. Furin is a cellular endoprotease. The green represents L domains which comprise the ligand binding site. Each L domain consists of a single-stranded right-hand beta-helix. Here is the cascade of events for signaling through EGFR: The transmembrane has ligand-dependent tyrosine kinase activity. Binding of the hormone EGF causes receptor dimerization bringing the intracellular kinase domains (yellow PK_Try_Ser_Thr) together activating them. When active, they can phosphorylate each other (autophosphorylation) or other proteins. When the receptor is autophosphorylated, other proteins can bind to the cytoplasmic domain of the receptor Tyr kinase where they are phosphorylated. The target substrates phosphorylated by the receptor Tyr kinases are proteins with a common 100 amino acid domain called SH for src homology, based on structural homology to another cytoplasmic protein, Src. Src is an intracellular Tyr kinase activated when it binds through 2 SH domains to an autophosphorylated receptor Tyr kinase. Specifically, the SH2 domain has been shown to bind tyrosine-phosphorylated peptides. These domains target proteins to the autophosphorylated receptor Tyr kinase. Many proteins involved in signal transduction have SH2 domains. Some of these proteins also have catalytic domains with kinase activity. Others have phosphatase, transcription factor. or scaffolding domains. Figure \(20\) shows the hormone-depended dimerization of RTKs, their autophosphorylation, and the recruitment of proteins with SH2 domains. It is easier to envision how a GPCR is activated by binding its target hormone than for RTKs. GPCRs are single-chain proteins that pass through the membrane using 7-transmembrane helices. RTKs have a single transmembrane helix. The dimeric form of the RTK has some additional flexibility in the short region between the extracellular and transmembrane domains, allowing for the conformational changes necessary for the activation of the intracellular kinase domains. The crystal structure of the full EGFR is not known given the difficulties in crystallizing membrane proteins that span the membrane with a single alpha helix. However, separate structures of the dimeric extracellular domain and the intracellular kinase domains are known. Figure \(21\) shows an interactive iCn3D model of the dimeric extracellular and transmembrane domains of the epidermal growth factor receptor (3NJP). The two EGFR are shown in blue and magenta. The two bound EGFs are shown in cyan. Figure \(22\) shows an interactive iCn3D model of a dimer of the intracellular dimeric EGFR kinase domains in complex with an ATP analog-peptide conjugate (2GS6) The two EGFR kinase domains are shown in cyan and magenta. The ATP analogs in each domain (spacefill) are thiophosphoric acid O-((adenosyl-phospho)phospho_-S-acetamidyldiester. The peptide substrates (green stick) are 13mers with a tyrosine (sticks, labeled Y, minus the OHs) connected to the ATP analog. We've introduced the essential cell membrane players in signal transduction. • GPCRs which are not enzymes but which activate the bound heterotrimer G prortein Gα subunit, which then can activate or inhibit the integral membrane protein adenylate cylase or activate the membrane-bound enyzme protein kinase C. These enzymes produce second messengers cAMP (adenylate cyclase) and diacylglycerol (DAG)and IP3 (phospholipase C) • Receptor tyrosine kinases, ligand-activated receptor kinases, which can,on ligand-induced dimerization, autophosphorylate themselves or other target protein in the cell. In the next chapter section we will explore the next downstream effects in signaling, mediated by the second messengers cAMP, DAG and IP3 and the substrates phosphorylated by the ligand-active receptor tyrosine kinases.

textbooks/bio/Biochemistry/Fundamentals_of_Biochemistry_(Jakubowski_and_Flatt)/Unit_IV_-_Special_Topics/28%3A_Biosignaling_-_Capstone_Volume_I/28.02%3A_At_the_cell_membrane-_receptors_and_receptor_enzymes.txt