Biosynthesis of Amino Acids, Nucleotides, and Related Molecules

CHAPTER 22 BIOSYNTHESIS OF AMINO ACIDS, NUCLEOTIDES, AND RELATED MOLECULES Amino acids and nucleotides are the precursors to proteins and nucleic acids, respectively. They also give rise to numerous neurotransmitters, metabolic cofactors, and other molecules of biological importance. The supply of biologically available nitrogen can be limiting in many environments. Atmospheric N2 is relatively inert and must be converted to other forms, such as ammonia and nitrate, to accommodate the requirements of life. A complex web of enzymatic processes, based primarily in microorganisms, interconverts the various molecular forms that comprise the global inventory of reactive nitrogen. Oxygen and nitrogen metabolism are interlinked. The oxidation and reduction of the various forms of nitrogen in the biosphere oen involve oxygen. Regulation again plays an important role. Many of the processes covered in this chapter are regulated to carefully conserve a critical and limited resource. Tight regulation is needed to maintain balanced supplies of amino acids and nucleotides. The metabolic flux through most of these pathways is far less than for carbohydrate or lipid biosynthetic pathways; most amino acids and nucleotides are not stored, but are synthesized as they are needed. The amino acids glutamate and glutamine represent the entry point where reactive forms of nitrogen are incorporated into biological systems. Reflecting their importance, the concentrations of these amino acids are sufficiently elevated in many tissues that they are major contributors to the electrochemical environment of cells. The prominent role of these two amino acids is a universal feature of animate nitrogen metabolism, yet another molecular manifestation of the shared evolutionary history of all organisms on the planet. Like other anabolic pathways, the reaction sequences in amino acid and nucleotide biosynthesis are endergonic and reductive. They use ATP as a source of metabolic energy and they use a reduced electron carrier (usually NADPH) as a reductant. Discussing the biosynthetic pathways for amino acids and nucleotides together is a sound approach, not only because both classes of molecules contain nitrogen, but also because the two sets of pathways are extensively intertwined, with several key intermediates in common. Certain amino acids or parts of amino acids are incorporated into the structure of purines and pyrimidines, and in one case, part of a purine ring is incorporated into an amino acid (histidine). The two sets of pathways also share much common chemistry, in particular a preponderance of reactions involving the transfer of nitrogen or one-carbon groups. The sheer number of steps and variety of intermediates in the pathways described here can be intimidating to the beginning biochemistry student. These pathways are best approached by maintaining a focus on metabolic principles we have already discussed, on key intermediates and precursors, and on common classes of reactions. Even a cursory look at the chemistry can be rewarding, for some of the most unusual chemical transformations in biological systems occur in these pathways; for instance, we find prominent examples of the rare biological use of the metals molybdenum, selenium, and vanadium. The effort also offers a practical dividend, especially for students of human or veterinary medicine. Many genetic diseases of humans and animals have been traced to an absence of one or more enzymes of amino acid and nucleotide metabolism, and many pharmaceuticals in common use to combat infectious diseases are inhibitors of enzymes in these pathways — as are a number of the most important agents in cancer chemotherapy. 22.1 Overview of Nitrogen Metabolism The biosynthetic pathways leading to amino acids and nucleotides share a requirement for nitrogen. Soluble, biologically useful nitrogen compounds are generally scarce in natural environments; thus, most organisms use ammonia, amino acids, and nucleotides economically. Available amino acids, purines, and pyrimidines formed during metabolic turnover of proteins and nucleic acids are oen salvaged and reused. We begin by examining the pathways by which nitrogen from the environment is introduced into biological systems. A Global Nitrogen Cycling Network Maintains a Pool of Biologically Available Nitrogen The movement of nitrogen through the biosphere has been viewed historically as a cycle. However, our evolving understanding of the complexity of nitrogenous interconversions makes it clear that nitrogen moves through a complex web, rather than in a neat cycle (Fig. 22-1). Earth’s atmosphere is four-fihs molecular nitrogen (N2). However, N2 is too unreactive to be of use to living organisms. Conversion of N2 to forms that can support life (NH3,NO− 2,NO− 3) is called nitrogen fixation. The reduction of N2 to NH3 plays such a central role in making N2 available that this one reaction is oen considered synonymous with nitrogen fixation. In the biosphere, the metabolic processes of countless species function interdependently to salvage and reuse biologically available nitrogen. Most of the key reactions are carried out by bacteria and archaea. FIGURE 22-1 The global nitrogen web. The total amount of nitrogen fixed annually in the biosphere exceeds 1011kg; industrial sources of fixed nitrogen are now nearly as great. Reactions are identified with the processes they are involved in by colored arrows (see key). The oxidation number of the N atom is indicated on the vertical axis. [Information from M. M. M. Kuypers et al., Nat. Rev. Microbiol. 16:263, 2018, Fig. 1.] The reduction of atmospheric nitrogen (N2) by nitrogen-fixing bacteria and archaea to yield ammonia (NH3orNH+ 4), provides a useful anchor for our discussion. This critical process, described in detail in Section 22.2, provides most of the reduced nitrogen for incorporation into biomolecules. Free ammonia does not build up, and reduction is balanced by oxidation. Bacteria that derive their energy by oxidizing ammonia to nitrite (NO− 2) and ultimately nitrate (NO− 3) are abundant and active in both terrestrial and marine environments. The processes of converting ammonia to nitric oxide, nitrite, and finally nitrate are known as nitrification (Fig. 22-1, purple arrows). Individual bacterial or archaeal species may promote one or more of these steps. Atmospheric N2 must be replaced in order to maintain levels at a steady-state concentration. Some of the replacement comes from reduction of nitrate and nitrite. The reduction of nitrate and nitrite to N2 under anaerobic conditions, a process called denitrification (Fig. 22-1, red arrows), is carried out by specialized microorganisms in all three domains of life. These organisms use NO− 3 or NO− 2 rather than O2 as the ultimate electron acceptor in a series of reactions that (like oxidative phosphorylation) generates a transmembrane proton gradient, which is used to synthesize ATP. These microorganisms exist in all anoxic environments where nitrate is present, including soils, marine sediments, and eutrophic marine zones. An alternative path back to atmospheric N2 is provided by a group of bacteria that promote anaerobic ammonia oxidation, or anammox (Fig. 22-1, blue arrows). Anammox converts ammonia and nitrite to N2. As much as 50% to 70% of the NH3-to-N2 conversion in the biosphere may occur through this pathway, which went undetected until the 1980s. The obligate anaerobes that promote anammox are fascinating in their own right and are providing some useful solutions to waste-treatment problems (Box 22-1). BOX 22-1 UNUSUAL LIFESTYLES OF THE OBSCURE BUT ABUNDANT Air-breathers that we are, we can easily overlook the bacteria and archaea that thrive in anaerobic environments. Although rarely featured in introductory biochemistry textbooks, these organisms constitute much of the biomass of this planet, and their contributions to the balance of carbon and nitrogen in the biosphere are essential to all forms of life. As detailed in earlier chapters, the energy used to maintain living systems relies on the generation of proton gradients across membranes. Electrons derived from a reduced substrate are made available to electron carriers in membranes and pass through a series of electron transfers to a final electron acceptor. As a byproduct of this process, protons are released on one side of the membrane, generating the transmembrane proton gradient. The proton gradient is used to synthesize ATP or to drive other energy-requiring processes. For all eukaryotes, the reduced substrate is generally a carbohydrate (glucose or pyruvate) or a fatty acid and the electron acceptor is oxygen. Many bacteria and archaea are much more versatile. In anaerobic environments such as marine and freshwater sediments, the variety of life strategies is extraordinary. Almost any available redox pair can be an energy source for some specialized organism or group of organisms. For example, a large number of lithotrophic bacteria (a lithotroph is a chemotroph that uses inorganic energy sources) have a hydrogenase that uses molecular hydrogen to reduce NAD +: H2+ NAD + hydrogenase −−−−−−→ NAD H + H+ The NADH is a source of electrons for a variety of membrane-bound electron acceptors, generating the proton gradient needed for ATP synthesis. Other lithotrophs oxidize sulfur compounds (H2S, elemental sulfur, or thiosulfate) or ferrous iron. A widespread group of archaea called methanogens, all strict anaerobes, extract energy from the reduction of CO2 to methane. And this is just a small sampling of what anaerobic organisms do for a living. Their metabolic pathways are replete with interesting reactions and highly specialized cofactors unknown in our own world of obligate aerobic metabolism. Study of these organisms can yield practical dividends. It can also provide clues about the origins of life on an early Earth, in an atmosphere that lacked molecular oxygen. The web of reactions that defines nitrogen utilization in the biosphere depends on a wide range of specialized bacteria. Some nitrifying bacteria oxidize ammonia to nitrites, and some oxidize the resulting nitrites to nitrates (see Fig. 22-1, purple arrows). Nitrate is second only to O2 as a biological electron acceptor, and a great many bacteria and archaea can catalyze the denitrification of nitrates and nitrites to nitrogen, which the nitrogen-fixing bacteria then convert back into ammonia. Ammonia is a major pollutant in sewage and in farm animal waste, and it is a byproduct of fertilizer manufacture and oil refining. Waste-treatment plants have generally made use of communities of nitrifying and denitrifying bacteria to convert ammonia waste to atmospheric nitrogen. The process is expensive, requiring inputs of organic carbon and oxygen. In the 1960s and 1970s, a few articles appeared in the research literature suggesting that ammonia could be oxidized to nitrogen anaerobically, using nitrite as an electron acceptor; this process was called anammox. The reports received little notice until bacteria promoting anammox were discovered in a waste-treatment system in Del , the Netherlands, in the mid-1980s. A team of Dutch microbiologists led by Gijs Kuenen and Mike Jetten began to study these bacteria, which were soon identified as belonging to an unusual bacterial phylum, Planctomycetes. Some surprises were to follow. The biochemistry underlying the anammox process was slowly unraveled (Fig. 1). Hydrazine (N2H4), a highly reactive molecule used as a rocket fuel, was an unexpected intermediate. As a small molecule, hydrazine is both highly toxic and difficult to contain. It readily diffuses across typical phospholipid membranes. The anammox bacteria solve this problem by sequestering hydrazine in a specialized organelle, dubbed the anammoxosome. The membrane of this organelle is composed of lipids known as ladderanes (Fig. 2), never before encountered in biology. The fused cyclobutane rings of ladderanes stack tightly to form a very dense barrier, greatly slowing the release of hydrazine. Cyclobutane rings, with their unusual bond angles, are strained and difficult to synthesize; the bacterial mechanisms for synthesizing these lipids are not yet known. FIGURE 1 The anammox reactions. Ammonia and hydroxylamine are converted to hydrazine and H2O by hydrazine hydrolase, and the hydrazine is oxidized by hydrazine-oxidizing enzyme, generating N2 and protons. The protons create a proton gradient for ATP synthesis. On the anammoxosome exterior, protons are used by the nitrite-reducing enzyme, producing hydroxylamine and completing the cycle. All of the anammox enzymes are embedded in the anammoxosome membrane. [Information from L. A. van Ni rik et al., FEMS Microbiol. Lett. 233:10, 2004, Fig. 4.] FIGURE 2 (a) Ladderane lipids of the anammoxosome membrane. The mechanism for synthesis of the unstable fused cyclobutane ring structures is unknown. (b) Ladderanes can stack to form a very dense, impermeable, hydrophobic membrane structure, allowing sequestration of the hydrazine produced in the anammox reactions. [Information from L. A. van Ni rik et al., FEMS Microbiol. Lett. 233:7, 2004, Fig. 3.] The anammoxosome was a surprising finding. Bacterial cells generally do not have compartments, and the lack of a membrane-enclosed nucleus is o en cited as the primary distinction between eukaryotes and bacteria. One type of organelle in a bacterium was interesting enough, but microbiologists also found that planctomycetes have a nucleus: their chromosomal DNA is contained within a membrane (Fig. 3). Planctomycetes are an ancient bacterial line with multiple genera, three of which are known to carry out the anammox reactions. Discovery of this subcellular organization has prompted further research to trace the origin of the planctomycetes and the evolution of eukaryotic nuclei. Further study of this group may ultimately bring us closer to a key goal of evolutionary biology: a description of the organism affectionately referred to as LUCA — the last universal common ancestor of all life on our planet. FIGURE 3 Transmission electron micrograph of a cross section through Gemmata obscuriglobus, showing the DNA in a nucleus (N) with enclosing nuclear envelope (NE). Bacteria of the Gemmata genus (phylum Planctomycetes) do not promote the anammox reactions. For now, the anammox bacteria offer a major advance in waste treatment, reducing the cost of ammonia removal by as much as 90% (the conventional denitrification steps are eliminated completely, and the aeration costs associated with nitrification are lower) and reducing the release of polluting byproducts. Clearly, a greater familiarity with the bacterial underpinnings of the biosphere can pay big dividends as we deal with the environmental challenges of the twenty-first century. Fixation of atmospheric N2 is not the only source of reduced ammonia for biological systems. Much of it comes from an alternative fate of nitrate that circumvents denitrification. More than 90% of the NH+4 generated by vascular plants, algae, and microorganisms comes from nitrate assimilation, a two-step reductive process that bypasses atmospheric N2. First NO− 3 is reduced to NO− 2 by nitrate reductase, then the NO− 2 is reduced to NH+4 in a six-electron transfer catalyzed by nitrite reductase (Fig. 22-2). Both reactions involve chains of electron carriers and cofactors we have not yet encountered. Nitrate reductase is a large, soluble protein (Mr220,000). Within the enzyme, a pair of electrons, donated by NADH, flows through — SH groups of cysteine, FAD, and a cytochrome (cytb557), then to a novel cofactor containing molybdenum, before reducing the substrate NO− 3 to NO− 2. FIGURE 22-2 Nitrate assimilation by nitrate reductase and nitrite reductase. (a) Nitrate reductases of plants and bacteria catalyze the two-electron reduction of NO−3 to NO−2, in which a novel Mo-containing cofactor plays a central role. NADH is the electron donor. (b) Nitrite reductase converts the product of nitrate reductase into NH+4 in a six-electron, eight-proton transfer process in which the metallic center in siroheme carries electrons and the carboxyl groups of siroheme may donate protons. The initial source of electrons is reduced ferredoxin. The nitrite reductase of plants is located in the chloroplasts and receives its electrons from ferredoxin (which is reduced in the light-dependent reactions of photosynthesis; see Section 20.2). Six electrons, donated one at a time by ferredoxin, pass through a 4Fe-4S center in the enzyme, then through a novel hemelike molecule (siroheme) before reducing NO− 2 to NH+4 (Fig. 22-2). Nonphotosynthetic microbes possess a distinct nitrite reductase for which NADPH is the electron donor. Human activity presents an increasing challenge to the global nitrogen balance, and to all life in the biosphere supported by that balance. Fixed nitrogen is increasingly necessary to boost production in agriculture. Industrial nitrogen-based fertilizers now contribute as much ammonia and other reactive nitrogen species to the biosphere as do natural processes. Nonfarming manufacturing activity releases additional reactive nitrogen into the atmosphere, including nitric oxide, a prominent greenhouse gas. Controlling the damaging effects of agricultural runoff and industrial pollutants will remain an important component of the continuing effort to expand the food supply for a growing human population. Nitrogen Is Fixed by Enzymes of the Nitrogenase Complex The availability of fixed nitrogen, an essential nutrient, may have limited the size of the primordial biosphere. As early cells acquired a capacity to fix atmospheric nitrogen, the biosphere expanded. Evidence for biological nitrogen fixation has been found in sedimentary rocks more than 3 billion years old. In the biosphere of today, only certain bacteria and archaea can fix atmospheric N2. These organisms, called diazotrophs, include the cyanobacteria of soils and fresh and salt waters, methanogenic archaea (strict anaerobes that obtain energy and carbon by converting H2 and CO2 to methane), other kinds of free-living soil bacteria such as Azotobacter species, and the nitrogen-fixing bacteria that live as symbionts in the root nodules of leguminous plants. The most important product of nitrogen fixation is ammonia, which can be used by all organisms either directly or aer its conversion to other soluble compounds such as nitrites, nitrates, or amino acids. The reduction of nitrogen to ammonia is an exergonic reaction: N2+ 3H2 →  2NH3 ΔG′°= −33.5 kJ /mol The N≡N triple bond, however, is very stable, with a bond energy of 930 kJ/mol. Nitrogen fixation therefore has an extremely high activation energy, and atmospheric nitrogen is almost chemically inert under normal conditions. Ammonia is produced industrially by the Haber process (named for its inventor, Fritz Haber), which requires temperatures of 400 to 500 °C and nitrogen and hydrogen at pressures of tens of thousands of kilopascals (several hundred atmospheres) to provide the necessary activation energy. Biological nitrogen fixation must occur at biological temperatures and at 0.8 atm of nitrogen, and the high activation barrier is overcome by other means. This is accomplished, at least in part, by the binding and hydrolysis of ATP. The overall reaction can be written N2+ 10H+ + 8e–+ 16AT P →  2NH+4 + 16AD P + 16Pi+ H2 Biological nitrogen fixation to produce ammonia is carried out by a highly conserved complex of proteins called the nitrogenase complex; its central components are dinitrogenase reductase and dinitrogenase (Fig. 22-3a). Dinitrogenase reductase (Mr60,000) is a dimer of two identical subunits. It contains a single 4Fe-4S redox center (see Fig. 19-5), bound between the subunits, and can be oxidized and reduced by one electron. It also has two binding sites for ATP/ADP (one site on each subunit). Dinitrogenase (Mr240,000), an α2β2 tetramer, has two Fe- containing cofactors that transfer electrons (Fig. 22-3b). One, the P cluster, has a pair of 4Fe-4S centers; these share a sulfur atom, making an 8Fe-7S center. The second cofactor in dinitrogenase, the FeMo cofactor, is a novel structure composed of 7 Fe atoms, 9 inorganic S atoms, a Cys side chain, and a single carbon atom in the center of the FeS cluster. Also part of the cofactor is a molybdenum atom, with ligands that include three inorganic S atoms, a His side chain, and two oxygen atoms from a molecule of homocitrate that is an intrinsic part of the FeMo cofactor. FIGURE 22-3 Enzymes and cofactors of the nitrogenase complex. (a) The holoenzyme consists of two identical dinitrogenase reductase molecules (green), each with a 4Fe-4S redox center and binding sites for two ATP, and two identical dinitrogenase heterodimers (purple and blue), each with a P cluster (Fe-S center) and an FeMo cofactor. In this structure, ADP is bound in the ATP site, to make the crystal more stable. (b) The electron- transfer cofactors. A P cluster is shown here in its reduced (top) and oxidized (middle) forms. The FeMo cofactor (bottom) has a Mo atom with three S ligands, a His ligand, and two oxygen ligands from a molecule of homocitrate. In some organisms, the Mo atom is replaced with a vanadium atom. (Fe is shown in orange, S in yellow.) [Data from (a) PDB ID 1N2C, H. Schindelin et al., Nature 387:370, 1997; (b) Pred: PDB ID 3MIN, and Pox: PDB ID 2MIN, J. W. Peters et al., Biochemistry 36:1181, 1997; FeMo cofactor: PDB ID 1M1N, O. Einsle et al., Science 297:1696, 2002.] There are two additional forms of nitrogenase. One includes a dinitrogenase with a vanadium-containing cofactor rather than molybdenum (VFe); the other contains a second Fe atom (FeFe). Each of the nitrogenase complexes is encoded by a separate set of genes. The FeMo nitrogenase complex is the ancestral type, and all nitrogen-fixing bacteria and archaea contain it. Some species can produce one or both of the alternative VFe or FeFe types. Although the alternative enzymes are somewhat less efficient, they may play important roles in environments in which molybdenum is limiting or absent. They may also permit some additional reactions to occur. The vanadium nitrogenase system of Azotobacter vinelandii has the remarkable capacity to catalyze the reduction of carbon monoxide (CO) to ethylene (C2H4), ethane, and propane. Nitrogen fixation to produce ammonia is carried out by a highly reduced form of dinitrogenase and requires eight electrons: six for the reduction of N2 and two to produce one molecule of H2. Production of H2 is an obligate part of the reaction mechanism, but the biological role of H2 in the process is not understood. Dinitrogenase is reduced by the transfer of electrons from dinitrogenase reductase (Fig. 22-4). The dinitrogenase tetramer has two binding sites for the reductase. The required eight electrons are transferred from reductase to dinitrogenase one at a time: a reduced reductase molecule binds to the dinitrogenase and transfers a single electron, then the oxidized reductase dissociates from dinitrogenase, in a repeating cycle. Each turn of the cycle requires the hydrolysis of two ATP molecules by the dimeric reductase. The immediate source of electrons to reduce dinitrogenase reductase varies, with reduced ferredoxin (see Section 20.2), reduced flavodoxin, and perhaps other sources playing a role. In at least one species, the ultimate source of electrons to reduce ferredoxin is pyruvate.

FIGURE 22-4 Electron path in nitrogen fixation by the nitrogenase complex. Electrons are transferred from pyruvate to dinitrogenase via ferredoxin (or flavodoxin) and dinitrogenase reductase. Dinitrogenase reductase reduces dinitrogenase one electron at a time, with at least six electrons required to fix one molecule of N2. Two additional electrons are used to reduce 2H+ to H2 in a process that obligatorily accompanies nitrogen fixation in anaerobes, making a total of eight electrons required per N2 molecule. The subunit structures and metal cofactors of the dinitrogenase reductase and dinitrogenase proteins are described in the text and in Figure 22-3. In the reaction carried out by dinitrogenase reductase, both ATP binding and ATP hydrolysis bring about protein conformational changes that help overcome the high activation energy of nitrogen fixation. The binding of two ATP molecules to the reductase shis the reduction potential (E ′°) of this protein from −300 to −420mV, an enhancement of its reducing power that is required to transfer electrons through dinitrogenase to N2; the standard reduction potential for the half-reaction N2+ 6H+ + 6e– →  2NH3 is −0.34V. The ATP molecules are then hydrolyzed just before the actual transfer of one electron to dinitrogenase. ATP binding and hydrolysis change the conformation of nitrogenase reductase in two regions, which are structurally homologous with the switch 1 and switch 2 regions of the GTP- binding proteins involved in biological signaling (see Fig. 12-12). ATP binding produces a conformational change that brings the 4Fe-4S center of the reductase closer to the P cluster of dinitrogenase (from 18 Å to 14 Å away), which facilitates electron transfer between the reductase and dinitrogenase. The details of electron transfer from the P cluster to the FeMo cofactor, and the means by which eight electrons are accumulated by nitrogenase, are not yet known in detail. Two pathways that conform to available data, both involving the Mo atom as a central player, are illustrated in Figure 22-5. FIGURE 22-5 Two reasonable hypotheses for the intermediates involved in N2 reduction. In both scenarios, the FeMo cofactor (abbreviated as M here) plays a central role, binding directly to one of the nitrogen atoms of N2 and remaining bound throughout the sequence of reduction steps. [Information from L. C. Seefeldt et al., Annu. Rev. Biochem. 78:701, 2009, Fig. 9.] The nitrogenase complex is remarkably unstable in the presence of oxygen. The reductase is inactivated in air, with a half-life of 30 seconds; dinitrogenase has a half-life of only 10 minutes in air. Free-living bacteria that fix nitrogen cope with this problem in a variety of ways. Some live only anaerobically or repress nitrogenase synthesis when oxygen is present. Some aerobic species, such as A. vinelandii, partially uncouple electron transfer from ATP synthesis so that oxygen is burned off as rapidly as it enters the cell (see Box 19-1). When fixing nitrogen, cultures of these bacteria increase in temperature as a result of their efforts to rid themselves of oxygen. The symbiotic relationship between leguminous plants and the nitrogen-fixing bacteria in their root nodules (Fig. 22-6) takes care of both the energy requirements and the oxygen lability of the nitrogenase complex. The energy required for nitrogen fixation was probably the evolutionary driving force for this plant-bacteria association. The bacteria in root nodules have access to a large reservoir of energy in the form of abundant carbohydrate and citric acid cycle intermediates made available by the plant. This may allow the bacteria to fix hundreds of times more nitrogen than do their free-living cousins under conditions generally encountered in soils. To solve the oxygen-toxicity problem, the bacteria in root nodules are bathed in a solution of the oxygen- binding heme protein leghemoglobin, produced by the plant (although the heme may be contributed by the bacteria). Leghemoglobin binds all available oxygen so that it cannot interfere with nitrogen fixation, and it efficiently delivers the oxygen to the bacterial electron-transfer system. The benefit to the plant, of course, is a ready supply of reduced nitrogen. In fact, the bacterial symbionts typically produce far more NH3 than is needed by their symbiotic partner; the excess is released into the soil. The efficiency of the symbiosis between plants and bacteria is evident in the enrichment of soil nitrogen brought about by leguminous plants. This enrichment of NH3 in the soil is the basis of crop rotation methods, in which plantings of nonleguminous plants (such as maize) that extract fixed nitrogen from the soil are alternated with plantings of legumes such as alfalfa, peas, or clover. FIGURE 22-6 Nitrogen-fixing nodules. (a) Pea plant (Pisum sativum) root nodules containing the nitrogen-fixing bacterium Rhizobium leguminosarum. The nodules are pink due to the presence of leghemoglobin; this heme protein has a very high binding affinity for oxygen, which strongly inhibits nitrogenase. (b) Artificially colorized electron micrograph of a thin section through a pea root nodule. Symbiotic nitrogen-fixing bacteria, or bacteroids (red), live inside the nodule cell, surrounded by the peribacteroid membrane (blue). Bacteroids produce the nitrogenase complex that converts atmospheric nitrogen (N2) to ammonium (NH+4); without the bacteroids, the plant is unable to utilize N2. (The cell nucleus is shown in yellow/green. Not visible in this micrograph are other organelles of the infected root cell that are normally found in plant cells.) Nitrogen fixation is energetically costly: 16 ATP and 8 electrons yield only 2 NH3. It is therefore not surprising that the process is tightly regulated so that NH3 is produced only when needed. High [ADP], an indicator of low [ATP], is a strong inhibitor of nitrogenase. NH+ 4 represses the expression of the ~20 nitrogen fixation (nif) genes, effectively shutting down the pathway. Covalent alteration of nitrogenase is also used in some diazotrophs to control nitrogen fixation in response to the availability of NH+ 4 in the surroundings. Transfer of an ADP- ribosyl group from NADH to a specific Arg residue in the nitrogenase reductase shuts down N2 fixation in Rhodospirillum, for example. This is the same covalent modification that we saw in the case of G protein inhibition by the toxins of cholera and pertussis (see Fig. 12-14). Nitrogen fixation is the subject of intense study because of its immense practical importance. Industrial production of ammonia for use in fertilizers requires a large and expensive input of energy, and this has spurred a drive to develop recombinant or transgenic organisms that can fix nitrogen. In principle, recombinant DNA techniques (Chapter 9) might be used to transfer the DNA that encodes the enzymes of nitrogen fixation into non-nitrogen-fixing bacteria and plants. However, those genes alone will not suffice. About 20 genes are essential to nitrogenase activity in bacteria, many of them needed for the synthesis, assembly, and insertion of the cofactors. There is also the problem of protecting the enzyme in its new setting from destruction by oxygen. In all, there are formidable challenges in engineering new nitrogen-fixing plants. Success in these efforts will depend on overcoming the problem of oxygen toxicity in any cell that produces nitrogenase. Ammonia Is Incorporated into Biomolecules through Glutamate and Glutamine Reduced nitrogen in the form of NH+4 is assimilated into amino acids and then into other nitrogen-containing biomolecules. Two amino acids, glutamate and glutamine, provide the critical entry point. Recall that these same two amino acids play central roles in the catabolism of ammonia and amino groups in amino acid oxidation (Chapter 18). Glutamate is the source of amino groups for most other amino acids, through transamination reactions (the reverse of the reaction shown in Fig. 18-4). The amide nitrogen of glutamine is a source of amino groups in a wide range of biosynthetic processes. In most types of cells, and in extracellular fluids in higher organisms, one or both of these amino acids are present at higher concentrations — sometimes an order of magnitude or more higher — than other amino acids. An Escherichia coli cell requires so much glutamate that this amino acid is one of the primary solutes in the cytosol. Its concentration is regulated not only in response to the cell’s nitrogen requirements but also to maintain an osmotic balance between the cytosol and the external medium. The biosynthetic pathways to glutamate and glutamine are simple, and all or some of the steps occur in most organisms. The most important pathway for the assimilation of NH+ 4 into glutamate requires two reactions. The net effect is to convert glutamate, α -ketoglutarate, and ammonia into two molecules of glutamate. First, NH+ 4 is reacted with glutamate to produce glutamine, using the enzyme glutamine synthetase. This reaction takes place in two steps, with enzyme-bound γ -glutamyl phosphate as an intermediate (see Fig. 18-8): (22-1) (1) Glutamate+ AT P → γ-glutamyl phosphate+ AD P (2) γ-Glutamyl phosphate+ NH+ 4 → glutamine+ Pi+ H+ Sum: Glutamate+ NH+4 + AT P → glutamine+ AD P + Pi+ H+ Glutamine synthetase is found in all organisms. In addition to its importance for NH+4 assimilation in bacteria, it has a central role in amino acid metabolism in mammals, converting free NH+4, which is toxic, to glutamine for transport in the blood (Chapter 18). In the second reaction needed for NH+ 4 assimilation, the glutamine reacts with α -ketoglutarate to generate two molecules of glutamate. In bacteria and plants, this reaction is catalyzed by glutamate synthase. (An alternative name for this enzyme, glutamate:oxoglutarate aminotransferase, yields the acronym GOGAT, by which the enzyme also is known.) α -Ketoglutarate, an intermediate of the citric acid cycle, undergoes reductive amination with glutamine as nitrogen donor: (22-2) The net reaction of glutamine synthetase and glutamate synthase (Eqns 22-1 and 22-2) is Glutamate synthase is not present in animals, which instead maintain high levels of glutamate by processes such as the α-Ketoglutarate+ glutamine+ NAD (P)H + H+ → 2 glutamate+ α-Ketoglutarate+ NH+ 4 + NAD (P)H + AT P → glutamate+ NAD transamination of α -ketoglutarate during amino acid catabolism. Plants possess a second alternative form of glutamate synthase that uses reduced ferredoxin rather than NADPH as a source of reducing electrons. Glutamate can also be formed in yet another, albeit minor, pathway: the reaction of α -ketoglutarate and NH+ 4 to form glutamate in one step. This is catalyzed by glutamate dehydrogenase, an enzyme present in all organisms. Reducing power is furnished by NADPH: We encountered this reaction in the catabolism of amino acids (see Fig. 18-7). In eukaryotic cells, glutamate dehydrogenase is located in the mitochondrial matrix. The reaction equilibrium favors the reactants, and the Km for NH+ 4 (∼1 mM ) is so high that the reaction is not important for NH+ 4 assimilation in mammals. (Recall that the glutamate dehydrogenase reaction, in reverse (see Fig. 18-10), is one source of NH+ 4 destined for the urea cycle.) In microorganisms and plants, concentrations of NH+ 4 high enough for the glutamate dehydrogenase reaction to make a significant contribution to glutamate levels generally occur only when NH3 is added artificially to the growth environment. In general, soil bacteria and plants rely on the two-enzyme pathway outlined above (Eqns 22-1, 22-2). α-Ketoglutarate+ NH+4 + NAD (P)H + H+ → glutamate+ NAD ( Glutamine Synthetase Is a Primary Regulatory Point in Nitrogen Metabolism There are three known classes of glutamine synthetases. The class I enzyme (GSI, found in bacteria) has 12 identical subunits of Mr50,000 (Fig. 22-7) and is regulated both allosterically and by covalent modification. The class II enzyme (GSII, found in eukaryotes and some bacteria) has 10 identical subunits. The third class of glutamine synthetases (GSIII), so far found only in two bacterial species, are much larger enzymes, consisting of a double-ringed dodecamer of identical chains.

FIGURE 22-7 Subunit structure of bacterial type I glutamine synthetase. This view shows 6 of the 12 identical subunits; a second layer of 6 subunits lies directly beneath those shown. Each of the 12 subunits has an active site, where ATP and glutamate are bound in orientations that favor transfer of a phosphoryl group from ATP to the side-chain carboxyl of glutamate. In this crystal structure, ADP occupies the ATP site. [Data from PDB ID 2GLS, M. M. Yamashita et al., J. Biol. Chem. 264:17,681, 1989.] Befitting their central metabolic role as an entry point for reduced nitrogen, GSI glutamine synthetases are highly regulated. In enteric bacteria such as E. coli, the regulation is unusually complex. Alanine, glycine, and at least six end products of glutamine metabolism are allosteric inhibitors (Fig. 22-8). Each inhibitor alone produces only partial inhibition, but the effects of multiple inhibitors are more than additive, and all eight together virtually shut down the enzyme. This is an example of cumulative feedback inhibition. This control mechanism provides a constant adjustment of glutamine levels to match immediate metabolic requirements. FIGURE 22-8 Allosteric regulation of glutamine synthetase. The enzyme undergoes cumulative regulation by six end products of glutamine metabolism. Alanine and glycine probably serve as indicators of the general status of amino acid metabolism in the cell. Superimposed on the allosteric regulation is inhibition by adenylylation of (addition of AMP to) T yr397, located near the enzyme’s active site (Fig. 22-9). This covalent modification increases sensitivity to the allosteric inhibitors, and activity decreases as more subunits are adenylylated. Both adenylylation and de-adenylylation are promoted by adenylyltransferase (AT in Fig. 22-9), part of a complex enzymatic cascade that responds to levels of glutamine, α -ketoglutarate, ATP, and Pi. The activity of adenylyltransferase is modulated by binding to a regulatory protein called PII, and the activity of PII, in turn, is regulated by covalent modification (uridylylation), again at a Tyr residue. The adenylyltransferase complex with uridylylated PII(PII-UMP) stimulates de-adenylylation, whereas the same complex with deuridylylated PII stimulates adenylylation of glutamine synthetase. Both uridylylation and deuridylylation of PII are brought about by a single enzyme, uridylyltransferase. Uridylylation is inhibited by binding of glutamine and Pi to uridylyltransferase and is stimulated by binding of α - ketoglutarate and ATP to PII. FIGURE 22-9 Second level of regulation of glutamine synthetase: covalent modifications. (a) An adenylylated Tyr residue. (b) Cascade leading to adenylylation (inactivation) of glutamine synthetase. AT represents adenylyltransferase; UT, uridylyltransferase. PII is a regulatory protein, itself regulated by uridylylation. The regulation does not stop there. The uridylylated PII also mediates the activation of transcription of the gene encoding glutamine synthetase, thus increasing the cellular concentration of the enzyme; the deuridylylated PII brings about a decrease in transcription of the same gene. This mechanism involves an interaction of PII with additional proteins involved in gene regulation, of a type described in Chapter 28. The net result of this elaborate system of controls is a decrease in glutamine synthetase activity when glutamine levels are high, and an increase in activity when glutamine levels are low and α - ketoglutarate and ATP (substrates for the synthetase reaction) are available. The multiple layers of regulation permit a sensitive response in which glutamine synthesis is tailored to cellular needs. Several Classes of Reactions Play Special Roles in the Biosynthesis of Amino Acids and Nucleotides The pathways described in this chapter include a variety of interesting chemical rearrangements. Several of these recur and deserve special note before we progress to the pathways themselves. These are (1) transamination reactions and other rearrangements promoted by enzymes containing pyridoxal phosphate; (2) transfer of one-carbon groups, with either tetrahydrofolate (usually at the — CHO and — CH2OH oxidation levels) or S-adenosylmethionine (at the — CH3 oxidation level) as cofactor; and (3) transfer of amino groups derived from the amide nitrogen of glutamine. Pyridoxal phosphate (PLP), tetrahydrofolate (H4 folate), and S-adenosylmethionine (adoMet) are described in some detail in Chapter 18 (see Figs. 18-6, 18-17, and 18-18). Here we focus on amino group transfer involving the amide nitrogen of glutamine. More than a dozen known biosynthetic reactions use glutamine as the major physiological source of amino groups, and most of these occur in the pathways outlined in this chapter. As a class, the enzymes catalyzing these reactions are called glutamine amidotransferases. All have two structural domains: one binding glutamine, the other binding the second substrate, which serves as amino group acceptor (Fig. 22-10). A conserved Cys residue in the glutamine-binding domain is believed to act as a nucleophile, cleaving the amide bond of glutamine and forming a covalent glutamyl-enzyme intermediate. The NH3 produced in this reaction is not released, but instead is transferred through an “ammonia channel” to a second active site, where it reacts with the second substrate to form the aminated product. The covalent intermediate is hydrolyzed to the free enzyme and glutamate. If the second substrate must be activated, the usual method is the use of ATP to generate an acyl phosphate intermediate (R— OX in Fig. 22-10, where X is a phosphoryl group). The enzyme glutaminase acts similarly but uses H2O as the second substrate, yielding NH+ 4 and glutamate (see Fig. 18-8).

MECHANISM FIGURE 22-10 Proposed mechanism for glutamine amidotransferases. Each enzyme has two domains. The glutamine-binding domain contains structural elements conserved among many of these enzymes, including a Cys residue required for activity. The NH3-acceptor (second-substrate) domain varies. Two types of amino acceptors are shown. X represents an activating group, typically a phosphoryl group derived from ATP, that facilitates displacement of a hydroxyl group from R— OH by NH3. SUMMARY 22.1 Overview of Nitrogen Metabolism The molecular nitrogen that makes up 80% of Earth’s atmosphere is unavailable to most living organisms until it is reduced. A complex web of reactions converts atmospheric N2 to biologically useful forms and maintains a global balance between them. Prominent species that are interconverted within this web include ammonia (NH3 or NH+ 4; most reduced), nitrite (NO− 2), and nitrate (NO− 3; most oxidized). Conversion of N2 to ammonia is fixation. Nitrification constitutes the steps converting ammonia to nitrate. Conversion of nitrate to N2 constitutes denitrification. The alternative conversion of nitrate to ammonia is nitrate assimilation. Fixation of N2 as NH3 is carried out by the nitrogenase complex, in a reaction that requires large investments of ATP and of reducing power. The nitrogenase complex is highly labile in the presence of O2, and it is subject to regulation by the supply of NH3 . In living systems, reduced nitrogen is incorporated first into amino acids and then into a variety of other biomolecules, including nucleotides. The key entry point is the amino acid glutamate. Glutamate and glutamine are the nitrogen donors in a wide range of biosynthetic reactions. Glutamine synthetase, which catalyzes the formation of glutamine from glutamate, is a main regulatory enzyme of nitrogen metabolism. The amino acid and nucleotide biosynthetic pathways make repeated use of the biological cofactors pyridoxal phosphate, tetrahydrofolate, and S-adenosylmethionine. Pyridoxal phosphate is required for transamination reactions involving glutamate and for other amino acid transformations. One-carbon transfers require S-adenosylmethionine and tetrahydrofolate. Glutamine amidotransferases catalyze reactions that incorporate nitrogen derived from the amide group of glutamine. 22.2 Biosynthesis of Amino Acids All amino acids are derived from intermediates in glycolysis, the citric acid cycle, or the pentose phosphate pathway (Fig. 22-11). Nitrogen enters these biosynthetic pathways by way of glutamate and glutamine. Some pathways are simple, others are not. Ten of the amino acids are just one or several steps removed from the common metabolite from which they are derived. The biosynthetic pathways for others, such as the aromatic amino acids, are more complex. FIGURE 22-11 Overview of amino acid biosynthesis. The carbon skeleton precursors derive from three sources: glycolysis (light red), the citric acid cycle (blue), and the pentose phosphate pathway (purple). Organisms Vary Greatly in Their Ability to Synthesize the 20 Common Amino Acids Whereas most bacteria and plants can synthesize all 20 amino acids, mammals can synthesize only about half of them — generally those with simple pathways. These are oen called nonessential amino acids (see Table 18-1). The label is somewhat misleading, however, because innate biosynthetic pathways oen do not provide enough of these amino acids to support optimal growth and health. The remaining amino acids, the essential amino acids, cannot be synthesized by most mammals and must be obtained from food. A few amino acids are conditionally essential in mammals, required at particular stages of development. Unless otherwise indicated, the pathways for the 20 common amino acids presented below are those operative in bacteria. A useful way to organize these biosynthetic pathways is to group them into six families corresponding to their metabolic precursors (Table 22-1), and we use this approach to structure the detailed descriptions that follow. In addition to these six precursors, there is a notable intermediate in several pathways of amino acid and nucleotide synthesis: 5-phosphoribosyl-1- pyrophosphate (PRPP): TABLE 22-1 Amino Acid Biosynthetic Families, Grouped by Metabolic Precursor α -Ketoglutarate Glutamate Glutamine Proline Arginine Pyruvate Alanine Valine Leucine Isoleucine 3-Phosphoglycerate Serine Glycine Cysteine Phosphoenolpyruvate and erythrose 4-phosphate Tryptophan Phenylalanine Tyrosine Oxaloacetate Aspartate Asparagine Isoleucine Methionine Threonine Lysine Ribose 5-phosphate Histidine Essential amino acids in mammals. Derived from phenylalanine in mammals. PRPP is synthesized from ribose 5-phosphate derived from the pentose phosphate pathway (see Fig. 14-31), in a reaction a a a a a b a a a a a b catalyzed by ribose phosphate pyrophosphokinase: This enzyme is allosterically regulated by many of the biomolecules for which PRPP is a precursor. α -Ketoglutarate Gives Rise to Glutamate, Glutamine, Proline, and Arginine We have already described the biosynthesis of glutamate and glutamine. Proline is a cyclized derivative of glutamate (Fig. 22- Ribose5-phosphate+ AT P → 5-phosphoribosyl-1-pyrophosphate 12). In the first step of proline synthesis, ATP reacts with the γ -carboxyl group of glutamate to form an acyl phosphate, which is reduced by NADPH or NADH to glutamate γ - semialdehyde. This intermediate undergoes rapid spontaneous cyclization and is then reduced further to yield proline. FIGURE 22-12 Biosynthesis of proline and arginine from glutamate in bacteria. All five carbon atoms of proline arise from glutamate. In many organisms, glutamate dehydrogenase is unusual in that it uses either NADH or NADPH as a cofactor. The same may be true of other enzymes in these pathways. The γ -semialdehyde in the proline pathway undergoes a rapid, reversible cyclization to Δ1-pyrroline-5-carboxylate (P5C), with the equilibrium favoring P5C formation. Cyclization is averted in the ornithine/arginine pathway by acetylation of the α -amino group of glutamate in the first step and removal of the acetyl group a er the transamination. This acetyl group is highlighted in yellow. Although some bacteria lack arginase and thus the complete urea cycle, they can synthesize arginine from ornithine in steps that parallel the mammalian urea cycle, with citrulline and argininosuccinate as intermediates (see Fig. 18-10). Here, and in subsequent figures in this chapter, the reaction arrows indicate the linear path to the final products, without considering the reversibility of individual steps. For example, the step of the pathway leading to arginine that is catalyzed by N- acetylglutamate dehydrogenase is chemically similar to the glyceraldehyde 3-phosphate dehydrogenase reaction in glycolysis (see Fig. 14-7) and is readily reversible. Arginine is synthesized from glutamate via ornithine and the urea cycle in animals (Chapter 18). In principle, ornithine could also be synthesized from glutamate γ -semialdehyde by transamination, but the spontaneous cyclization of the semialdehyde in the proline pathway precludes a sufficient supply of this intermediate for ornithine synthesis. Bacteria have a de novo biosynthetic pathway for ornithine (and thus arginine) that parallels some steps of the proline pathway but includes two additional steps that avoid the problem of the spontaneous cyclization of glutamate γ -semialdehyde (Fig. 22-12). In the first step, the α -amino group of glutamate is blocked by an acetylation requiring acetyl-CoA; then, aer the transamination step, the acetyl group is removed to yield ornithine. The pathways to proline and arginine are somewhat different in mammals. Proline can be synthesized by the pathway shown in Figure 22-12, but it is also formed from arginine obtained from dietary or tissue protein. Arginase, a urea cycle enzyme, converts arginine to ornithine and urea (see Figs. 18-10, 18-26). The ornithine is converted to glutamate γ -semialdehyde by the enzyme ornithine γ -aminotransferase (Fig. 22-13). The semialdehyde cyclizes to Δ1-pyrroline-5-carboxylate, which is then converted to proline (Fig. 22-12). The pathway for arginine synthesis shown in Figure 22-12 is absent in mammals. When arginine from dietary intake or protein turnover is insufficient for protein synthesis, the ornithine δ - aminotransferase reaction operates in the direction of ornithine formation. Ornithine is then converted to citrulline and arginine in the urea cycle. FIGURE 22-13 Ornithine δ -aminotransferase reaction: a step in the mammalian pathway to proline. This enzyme is found in the mitochondrial matrix of most tissues. Although the equilibrium favors P5C formation, the reverse reaction is the only mammalian pathway for synthesis of ornithine (and thus arginine) when arginine levels are insufficient for protein synthesis. Serine, Glycine, and Cysteine Are Derived from 3-Phosphoglycerate The major pathway for the formation of serine is the same in all organisms (Fig. 22-14). In the first step, the hydroxyl group of 3- phosphoglycerate is oxidized by a dehydrogenase (using NAD +) to yield 3-phosphohydroxypyruvate. Transamination from glutamate yields 3-phosphoserine, which is hydrolyzed to free serine by phosphoserine phosphatase.

FIGURE 22-14 Biosynthesis of serine from 3-phosphoglycerate and of glycine from serine in all organisms. As indicated in the text, this is only one of multiple pathways to synthesize glycine. Serine (three carbons) is the precursor of glycine (two carbons) through removal of a carbon atom by serine hydroxymethyltransferase (Fig. 22-14). Tetrahydrofolate accepts the β carbon (C-3) of serine, which forms a methylene bridge between N-5 and N-10 to yield N5,N10-methylenetetrahydrofolate (see Fig. 18-17). The overall reaction, which is reversible, also requires pyridoxal phosphate. In the liver of vertebrates, glycine can be made by another route: the reverse of the reaction shown in Figure 18-20c, catalyzed by glycine synthase (also called glycine cleavage enzyme): CO2+ NH+4 + N5,N10-methylenetetrahydrofolate+ NAD H + H+ → glycine+ tetrahydrofolate+ NAD + Plants and bacteria produce the reduced sulfur required for the synthesis of cysteine (and methionine, described later) from environmental sulfates; the pathway is shown on the right side of Figure 22-15. Sulfate is activated in two steps to produce 3′- phosphoadenosine 5′-phosphosulfate (PAPS), which undergoes an eight-electron reduction to sulfide. The sulfide is then used in the formation of cysteine from serine in a two-step pathway. Mammals synthesize cysteine from two amino acids: methionine furnishes the sulfur atom, and serine furnishes the carbon skeleton. Methionine is first converted to S-adenosylmethionine (see Fig. 18-18), which can lose its methyl group to any of a number of acceptors to form S-adenosylhomocysteine (adoHcy). This demethylated product is hydrolyzed to free homocysteine, which undergoes a reaction with serine, catalyzed by cystathionine β -synthase, to yield cystathionine (Fig. 22-16). Finally, cystathionine δ -lyase, a PLP-requiring enzyme, catalyzes removal of ammonia and cleavage of cystathionine to yield free cysteine. FIGURE 22-15 Biosynthesis of cysteine from serine in bacteria and plants. The origin of reduced sulfur is shown in the pathway on the right. FIGURE 22-16 Biosynthesis of cysteine from homocysteine and serine in mammals. The homocysteine is formed from methionine. Three Nonessential and Six Essential Amino Acids Are Synthesized from Oxaloacetate and Pyruvate Alanine and aspartate are synthesized from pyruvate and oxaloacetate, respectively, by transamination from glutamate. Asparagine is synthesized by amidation of aspartate, catalyzed by the enzyme asparagine synthetase. The NH+4 is donated by glutamine. These are nonessential amino acids, and their simple biosynthetic pathways occur in all organisms. The malignant lymphocytes present in childhood acute lymphoblastic leukemia (ALL) produce little or no asparagine synthetase, and they are thus sensitive to asparagine depletion. The chemotherapy for ALL is administered together with an L- asparaginase derived from bacteria, with the enzyme functioning to reduce serum asparagine. The combined treatment results in a greater than 95% remission rate in cases of childhood ALL (L- asparaginase treatment alone produces remission in 40% to 60% of cases). However, the asparaginase treatment has some deleterious side effects, and about 10% of patients who achieve remission eventually suffer relapse, with tumors resistant to drug therapy. Researchers are now developing inhibitors of human asparagine synthetase to augment these therapies for childhood ALL. Methionine, threonine, lysine, isoleucine, valine, and leucine are essential amino acids; humans cannot synthesize them. Their biosynthetic pathways in bacteria are complex and interconnected (Fig. 22-17). In some cases, the pathways in bacteria, fungi, and plants differ significantly. FIGURE 22-17 Biosynthesis of six essential amino acids from oxaloacetate and pyruvate in bacteria: methionine, threonine, lysine, isoleucine, valine, and leucine. Some of the most complex pathways for amino acid biosynthesis are found here. Pathways are abbreviated to emphasize precursors and pathway products. Aspartate gives rise to methionine, threonine, and lysine. Branch points occur at aspartate β -semialdehyde, an intermediate in all three pathways, and at homoserine, a precursor of threonine and methionine. Threonine, in turn, is one of the precursors of isoleucine. The valine and isoleucine pathways share four enzymes (Fig. 22-17). Pyruvate gives rise to valine and isoleucine in pathways that begin with condensation of two carbons of pyruvate (in the form of hydroxyethyl thiamine pyrophosphate; see Fig. 14-13b) with another molecule of pyruvate (the valine path) or with α -ketobutyrate (the isoleucine path). The α -ketobutyrate is derived from threonine in a reaction that requires pyridoxal phosphate. An intermediate in the valine pathway, α -ketoisovalerate, is the starting point for a four-step branch pathway leading to leucine. Chorismate Is a Key Intermediate in the Synthesis of Tryptophan, Phenylalanine, and Tyrosine The three amino acid side chains that contain aromatic rings, tryptophan, phenylalanine, and tyrosine, present a special chemical problem for biosynthesis. Aromatic rings are not readily available in the environment, even though the benzene ring is very stable. The branched pathway to tryptophan, phenylalanine, and tyrosine, occurring in bacteria, fungi, and plants, is the main biological route of aromatic ring formation. It proceeds through ring closure of an aliphatic precursor followed by stepwise addition of double bonds. The first four steps produce shikimate, a seven-carbon molecule derived from erythrose 4-phosphate and phosphoenolpyruvate (Fig. 22-18). Shikimate is converted to chorismate in three steps that include the addition of three more carbons from another molecule of phosphoenolpyruvate. Chorismate is the first branch point of the pathway, with one branch leading to tryptophan, the other to phenylalanine and tyrosine. FIGURE 22-18 Biosynthesis of chorismate, an intermediate in the synthesis of aromatic amino acids in bacteria and plants. All carbons are derived from either erythrose 4-phosphate (light purple) or phosphoenolpyruvate (light red). Note that the NAD + required as a cofactor in step is released unchanged; it may be transiently reduced to NADH during the reaction, with formation of an oxidized reaction intermediate. Step is competitively inhibited by glyphosate (–COO— CH2— NH— CH2— PO2−3 ), the active ingredient in the widely used herbicide Roundup. The herbicide is relatively nontoxic to mammals, which lack this biosynthetic pathway. The intermediates quinate and shikimate are named a er the plants in which they have been found to accumulate. In the tryptophan branch (Fig. 22-19), chorismate is converted to anthranilate in a reaction in which glutamine donates the nitrogen that will become part of the indole ring. Anthranilate then condenses with PRPP. The indole ring of tryptophan is derived from the ring carbons and amino group of anthranilate plus two carbons derived from PRPP. The final reaction in the sequence is catalyzed by tryptophan synthase. This enzyme has an α2β2 subunit structure and can be dissociated into two α subunits and a β2 unit that catalyze different parts of the overall reaction: Indole-3-glycerolphosphate−−−−→αsubunitindole+ glyceraldehyde 3-phos Indole+ serine−−−−−→β2subunittryptophan+ H2O FIGURE 22-19 Biosynthesis of tryptophan from chorismate in bacteria and plants. In E. coli, enzymes catalyzing steps and are subunits of a single complex. The second part of the reaction requires pyridoxal phosphate (Fig. 22-20). Indole formed in the first part is not released by the enzyme, but instead moves through a channel from the α - subunit active site to one of the β -subunit active sites, where it condenses with a Schiff base intermediate derived from serine and PLP. Intermediate channeling of this type may be a feature of the entire pathway from chorismate to tryptophan. Enzyme active sites catalyzing different steps (sometimes not sequential steps) of the pathway to tryptophan are found on single polypeptides in some species of fungi and bacteria, but they are separate proteins in other species. In addition, the activity of some of these enzymes requires a noncovalent association with other enzymes of the pathway. These observations suggest that all the pathway enzymes are components of a large, multienzyme complex, a metabolon, in both bacteria and eukaryotes. Such complexes are generally not preserved intact when the enzymes are isolated using traditional biochemical methods (see Section 16.4). MECHANISM FIGURE 22-20 Tryptophan synthase reaction. (a) This enzyme catalyzes a multistep reaction with several types of chemical rearrangements. The PLP-facilitated transformations occur at the β carbon (C-3) of the amino acid, as opposed to the α - carbon reactions described in Figure 18-6. The β carbon of serine is attached to the indole ring system. (b) Indole generated on the α subunit (white) moves through a tunnel to the β subunit (blue), where it condenses with the serine moiety. [(b) Data from PDB ID 1KFJ, V. Kulik et al., J. Mol. Biol. 324:677, 2002.] In plants and bacteria, phenylalanine and tyrosine are synthesized from chorismate in pathways much less complex than the tryptophan pathway. The common intermediate is prephenate (Fig. 22-21). The final step in both cases is transamination with glutamate.

FIGURE 22-21 Biosynthesis of phenylalanine and tyrosine from chorismate in bacteria and plants. Conversion of chorismate to prephenate is a rare biological example of a Claisen rearrangement. Animals can produce tyrosine directly from phenylalanine through hydroxylation at C-4 of the phenyl group by phenylalanine hydroxylase; this enzyme also participates in the degradation of phenylalanine (see Figs. 18-23, 18-24). Tyrosine is considered a conditionally essential amino acid, or as nonessential insofar as it can be synthesized from the essential amino acid phenylalanine. Histidine Biosynthesis Uses Precursors of Purine Biosynthesis

The pathway to histidine in all plants and bacteria differs in several respects from other amino acid biosynthetic pathways. Histidine is derived from three precursors (Fig. 22-22): PRPP contributes five carbons, the purine ring of ATP contributes a nitrogen and a carbon, and glutamine supplies the second ring nitrogen. The key steps are condensation of ATP and PRPP, in which N-1 of the purine ring is linked to the activated C-1 of the ribose of PRPP (step in Fig. 22-22); purine ring opening that ultimately leaves N-1 and C-2 of adenine linked to the ribose (step ); and formation of the imidazole ring, a reaction in which glutamine donates a nitrogen (step ). The use of ATP as a metabolite rather than a high-energy cofactor is unusual — but not wasteful, because it dovetails with the purine biosynthetic pathway. The remnant of ATP that is released aer the transfer of N-1 and C-2 is 5-aminoimidazole-4-carboxamide ribonucleotide (AICAR), an intermediate of purine biosynthesis (see Fig. 22-35) that is rapidly recycled to ATP. FIGURE 22-22 Biosynthesis of histidine in bacteria and plants. Atoms derived from PRPP and ATP are shaded light red and blue, respectively. Two of the histidine nitrogens are derived from glutamine and glutamate (green). Note that the derivative of ATP remaining a er step (AICAR) is an intermediate in purine biosynthesis (see Fig. 22-35, step ), so ATP is rapidly regenerated. Amino Acid Biosynthesis Is under Allosteric Regulation As detailed in Chapter 13, the control of flux through a metabolic pathway oen reflects the activity of multiple enzymes in that pathway. In the case of amino acid synthesis, regulation oen takes place in part through feedback inhibition of the first reaction by the end product of the pathway. This first reaction is oen catalyzed by an allosteric enzyme that plays an important role in the overall control of flux through that pathway. As an example, Figure 22-23 shows the allosteric regulation of isoleucine synthesis from threonine. The end product, isoleucine, is an allosteric inhibitor of the first reaction in the sequence. In bacteria, such allosteric modulation of amino acid synthesis contributes to the minute-to-minute adjustment of pathway activity to cellular needs. FIGURE 22-23 Allosteric regulation of isoleucine biosynthesis. The first reaction in the pathway from threonine to isoleucine is inhibited by the end product, isoleucine. This was one of the first examples of allosteric feedback inhibition to be discovered. Allosteric regulation of an individual enzyme can be considerably more complex. An example is the remarkable set of allosteric controls exerted on glutamine synthetase of E. coli (Fig. 22-8). Six products derived from glutamine serve as negative feedback modulators of the enzyme, and the overall effects of these and other modulators are more than additive. Such regulation is called concerted inhibition. Additional mechanisms contribute to the regulation of the amino acid biosynthetic pathways. Because the 20 common amino acids must be made in the correct proportions for protein synthesis, cells have developed ways not only of controlling the rate of synthesis of individual amino acids but also of coordinating their formation. Such coordination is especially well developed in fast-growing bacterial cells. Figure 22-24 shows how E. coli cells coordinate the synthesis of lysine, methionine, threonine, and isoleucine, all made from aspartate. Several important types of inhibition patterns are evident. The step from aspartate to aspartyl-β -phosphate is catalyzed by three isozymes, each independently controlled by different modulators. This enzyme multiplicity prevents one biosynthetic end product from shutting down key steps in a pathway when other products of the same pathway are required. The steps from aspartate β - semialdehyde to homoserine and from threonine to α - ketobutyrate (detailed in Fig. 22-17) are also catalyzed by dual, independently controlled isozymes. One isozyme for the conversion of aspartate to aspartyl-β -phosphate is allosterically inhibited by two different modulators, lysine and isoleucine, whose action is more than additive — another example of concerted inhibition. The sequence from aspartate to isoleucine undergoes multiple, overlapping negative feedback inhibitions; for example, isoleucine inhibits the conversion of threonine to α - ketobutyrate (as described above), and threonine inhibits its own formation at three points: from homoserine, from aspartate β - semialdehyde, and from aspartate (see Fig. 22-17). This overall regulatory mechanism is called sequential feedback inhibition.

FIGURE 22-24 Interlocking regulatory mechanisms in the biosynthesis of several amino acids derived from aspartate in E. coli. Three enzymes (A, B, C) have either two or three isozyme forms, indicated by numerical subscripts. In each case, one isozyme (A2,B1,andC2) has no allosteric regulation; these isozymes are regulated by changes in the amount of enzyme synthesized. Synthesis of isozymes A2 and B1 is repressed when methionine levels are high, and synthesis of isozyme C2 is repressed when isoleucine levels are high. Enzyme A is aspartokinase; B, homoserine dehydrogenase; C, threonine dehydratase. Similar patterns are evident in the pathways leading to the aromatic amino acids. The first step of the early pathway to the common intermediate chorismate is catalyzed by the enzyme 2- keto-3-deoxy-D1-arabinoheptulosonate 7-phosphate (DAHP) synthase ( in Fig. 22-18). Most microorganisms and plants have three DAHP synthase isozymes. One is allosterically inhibited (feedback inhibition) by phenylalanine, another by tyrosine, and the third by tryptophan. This scheme helps the overall pathway to respond to cellular requirements for one or more of the aromatic amino acids. Additional regulation takes place aer the pathway branches at chorismate. For example, the enzymes catalyzing the first two steps of the tryptophan branch are subject to allosteric inhibition by tryptophan. SUMMARY 22.2 Biosynthesis of Amino Acids Plants and bacteria synthesize all 20 common amino acids. Mammals can synthesize about half; the others are required in the diet (essential or conditionally essential amino acids). Glutamate is formed by reductive amination of α - ketoglutarate and serves as the precursor of glutamine, proline, and arginine. The carbon chain of serine is derived from 3-phosphoglycerate. Serine is a precursor of glycine; the β -carbon atom of serine is transferred to tetrahydrofolate. In microorganisms, cysteine is produced from serine and from sulfide produced by the reduction of environmental sulfate. Mammals produce cysteine from methionine and serine by a series of reactions requiring S- adenosylmethionine and cystathionine. Alanine and aspartate (and thus asparagine) are formed from pyruvate and oxaloacetate, respectively, by transamination. Pyruvate and oxaloacetate also give rise to methionine, threonine, lysine, valine, isoleucine, and leucine via longer pathways. The aromatic amino acids (phenylalanine, tyrosine, and tryptophan) form by a pathway in which chorismate occupies a key branch point. Tyrosine can also be formed by hydroxylation of phenylalanine (and thus is considered conditionally essential). Phosphoribosyl pyrophosphate is a precursor of tryptophan and histidine. The pathway to histidine is interconnected with the purine synthetic pathway. The amino acid biosynthetic pathways are subject to allosteric end-product inhibition; the regulatory enzyme is usually the first in the sequence. Regulation of the various synthetic pathways is coordinated. 22.3 Molecules Derived from Amino Acids In addition to their role as the building blocks of proteins, amino acids are precursors of many specialized biomolecules, including hormones, coenzymes, nucleotides, alkaloids, cell wall polymers, porphyrins, antibiotics, pigments, and neurotransmitters. We describe here the pathways to a number of these amino acid derivatives. Glycine Is a Precursor of Porphyrins The biosynthesis of porphyrins, for which glycine is a major precursor, is our first example because of the central importance of the porphyrin nucleus in heme proteins such as hemoglobin and the cytochromes. The porphyrins are constructed from four molecules of the monopyrrole derivative porphobilinogen, which itself is derived from two molecules of δ -aminolevulinate. There are two major pathways to δ -aminolevulinate. In higher eukaryotes (Fig. 22-25a), glycine reacts with succinyl-CoA in the first step to yield α -amino-β -ketoadipate, which is then decarboxylated to δ -aminolevulinate. In plants, algae, and most bacteria, δ -aminolevulinate is formed from glutamate (Fig. 22- 25b). The glutamate is first esterified to glutamyl-tRNAGlu; reduction by NADPH converts the glutamate to glutamate 1- semialdehyde, which is cleaved from the tRNA. An aminotransferase converts the glutamate 1-semialdehyde to δ - aminolevulinate. FIGURE 22-25 Biosynthesis of δ -aminolevulinate. (a) In most animals, including mammals, δ -aminolevulinate is synthesized from glycine and succinyl-CoA. The atoms furnished by glycine are shown in red. (b) In bacteria and plants, the precursor of δ - aminolevulinate is glutamate. In all organisms, two molecules of δ -aminolevulinate condense to form porphobilinogen and, through a series of complex enzymatic reactions, four molecules of porphobilinogen come together to form protoporphyrin (Fig. 22-26). The iron atom is incorporated aer the protoporphyrin has been assembled, in a step catalyzed by ferrochelatase. Porphyrin biosynthesis is regulated in higher eukaryotes by heme, which serves as a feedback inhibitor of early steps in the synthetic pathway. Genetic defects in the biosynthesis of porphyrins can lead to the accumulation of pathway intermediates, causing a variety of human diseases known collectively as porphyrias (Box 22-2). FIGURE 22-26 Biosynthesis of heme from δ -aminolevulinate. Ac represents acetyl (— CH2COO−); Pr, propionyl (— CH2CH2COO−). BOX 22-2 MEDICINE On Kings and Vampires Porphyrias are a group of genetic diseases that result from defects in enzymes of the biosynthetic pathway from glycine to porphyrins (Fig. 1); specific porphyrin precursors accumulate in erythrocytes, body fluids, and the liver. The most common form is acute intermittent porphyria. Most individuals inheriting this condition are heterozygotes and are usually asymptomatic, because the single copy of the normal gene provides a sufficient level of enzyme function. However, certain nutritional or environmental factors (as yet poorly understood) can cause a buildup of δ -aminolevulinate and porphobilinogen, leading to attacks of acute abdominal pain and neurological dysfunction. King George III, British monarch during the American Revolution, suffered several episodes of apparent madness that tarnished the record of this otherwise accomplished man. The symptoms of his condition suggest that George III suffered from acute intermittent porphyria. FIGURE 1 The key to Figure 22-26 identifies the defective enzyme at each step. About 5,000 to 10,000 humans worldwide suffer from a genetic condition in which the activity of ferrochelatase (step ) is reduced, leading to abnormally high concentrations of protoporphyrin in tissues. When exposed to light, the excess protoporphyrin releases free radicals that damage cellular macromolecules, including proteins and lipids in cell membranes. The resulting cell damage and inflammation in the endothelial (skin) cells can be very painful. Patients are forced to adopt a lifestyle in which they avoid sunlight and even bright indoor light. A new drug called afamelanotide, slowly released over several months from a small implant under the skin, has shown success in alleviating the symptoms of sunlight exposure. The drug interacts with the melanocortin-1 receptor, stimulating melanin production and modulating the expression of antioxidants that eliminate the free radical species. Afamelanotide may also prove useful in treating vitiligo, a more common condition in which melanin production is not uniform and skin color is lost in blotches. One of the rarer porphyrias results in an accumulation of uroporphyrinogen I, an abnormal isomer of a protoporphyrin precursor. This compound stains the urine red, causes the teeth to fluoresce strongly in ultraviolet light, and makes the skin abnormally sensitive to sunlight. Many individuals with this porphyria are anemic because insufficient heme is synthesized. This genetic condition may have given rise to the vampire stories of folk legend. The symptoms of many porphyrias are now readily controlled with dietary changes or the administration of heme or heme derivatives. Heme Degradation Has Multiple Functions The iron-porphyrin (heme) group of hemoglobin, released from dying erythrocytes in the spleen, is degraded to yield free Fe2+ and, ultimately, bilirubin. The pathway also contributes the pigment present in mixtures of the bile salts derived from cholesterol. The first step in the two-step pathway to bilirubin, catalyzed by heme oxygenase, converts heme to biliverdin, a linear (open) tetrapyrrole derivative (Fig. 22-27 on p. 820). The other products of the reaction are free Fe2+ and CO. The Fe2+ is quickly bound by ferritin. Carbon monoxide is a poison that binds to hemoglobin (see Box 5-1), and the production of CO by heme oxygenase ensures that, even in the absence of environmental exposure, about 1% of an individual’s heme is complexed with CO. FIGURE 22-27 Bilirubin and its breakdown products. M represents methyl; V, vinyl; Pr, propionyl; E, ethyl. For ease of comparison, these structures are shown in linear form, rather than in their correct stereochemical conformations. Biliverdin is converted to bilirubin in the second step, catalyzed by biliverdin reductase. You can monitor the reactions in the breakdown of heme (Fig. 22-27) colorimetrically in a familiar in situ experiment. When you are bruised, the black and/or purple color results from hemoglobin released from damaged erythrocytes. Over time, the color changes to the green of biliverdin, and then to the yellow of bilirubin. Bilirubin is largely insoluble, and it travels in the bloodstream as one of many metabolites, fatty acids and others, complexed with serum albumin (see Fig. 17-3). In the liver, bilirubin is transformed to the bile pigment bilirubin diglucuronide. This product is sufficiently water-soluble to be secreted with other components of bile into the small intestine, where microbial enzymes convert it to several products, predominantly urobilinogen. Some urobilinogen is reabsorbed into the blood and transported to the kidney, where it is converted to urobilin, the compound that gives urine its yellow color. Urobilinogen remaining in the intestine is converted (in another microbe-dependent reaction) to stercobilin, which imparts the red-brown color to feces. Impaired liver function or blocked bile secretion causes bilirubin to leak from the liver into the blood, resulting in a yellowing of the skin and eyeballs, a condition called jaundice. In cases of jaundice, determination of the concentration of bilirubin in the blood may be useful in the diagnosis of underlying liver disease. Newborn infants sometimes develop jaundice because they have not yet produced enough glucuronyl bilirubin transferase to process their bilirubin. A traditional treatment to reduce excess bilirubin, exposure to a fluorescent lamp, causes a photochemical conversion of bilirubin to compounds that are more soluble and easily excreted. These pathways of heme breakdown play significant roles in protecting cells from oxidative damage and in regulating certain cellular functions. The CO produced by heme oxygenase is toxic at high concentrations, but at the very low concentrations generated during heme degradation it seems to have some regulatory and/or signaling functions. It acts as a vasodilator, much the same as (but less potent than) nitric oxide (discussed below). Low levels of CO also have some regulatory effects on neurotransmission. Bilirubin is the most abundant antioxidant in mammalian tissues and is responsible for most of the antioxidant activity in serum. Its protective effects seem to be especially important in the developing brain of newborn infants. The cell toxicity associated with jaundice may be due to bilirubin levels in excess of the serum albumin needed to solubilize it. Given these varied roles of heme degradation products, the degradative pathway is subject to regulation, mainly at the first step. Humans have at least three isozymes of heme oxygenase (HO). HO-1 is highly regulated; the expression of its gene is induced by a wide range of stress conditions, including shear stress, uncontrolled angiogenesis (development of blood vessels), hypoxia, hyperoxia, heat shock, exposure to ultraviolet light, hydrogen peroxide, and many other metabolic insults. HO-2 is found mainly in the brain and the testes, where it is continuously expressed. The third isozyme, HO-3, is not catalytically active, but may play a role in oxygen sensing. Amino Acids Are Precursors of Creatine and Glutathione Phosphocreatine, derived from creatine, is an important energy buffer in skeletal muscle (see Box 23-1). Creatine is synthesized from glycine and arginine (Fig. 22-28); methionine, in the form of S-adenosylmethionine, acts as methyl group donor.

FIGURE 22-28 Biosynthesis of creatine and phosphocreatine. Creatine is made from three amino acids: glycine, arginine, and methionine. This pathway shows the versatility of amino acids as precursors of other nitrogenous biomolecules. Glutathione (GSH), present in plants, animals, and some bacteria, oen at high levels, can be thought of as a redox buffer. It is derived from glutamate, cysteine, and glycine (Fig. 22-29). The γ -carboxyl group of glutamate is activated by ATP to form an acyl phosphate intermediate, which is then attacked by the α -amino group of cysteine. A second condensation reaction follows, with the α -carboxyl group of cysteine activated to an acyl phosphate to permit reaction with glycine. The oxidized form of glutathione (glutathione disulfide, or GSSG), produced in the course of its redox activities, contains two glutathione molecules linked by a disulfide bond. FIGURE 22-29 Glutathione metabolism. (a) Biosynthesis of glutathione. (b) Oxidized form of glutathione. Glutathione helps maintain the sulfhydryl groups of proteins in the reduced state and the iron of heme in the ferrous (Fe2+) state, and it serves as a reducing agent for glutaredoxin in deoxyribonucleotide synthesis (see Fig. 22-41). Its redox function is also used to remove toxic peroxides formed in the normal course of growth and metabolism under aerobic conditions: 2GSH + R— O— O— H →  GSSG + H2O + R— OH This reaction is catalyzed by glutathione peroxidase, a remarkable enzyme in that it contains a covalently bound selenium (Se) atom in the form of selenocysteine (see Fig. 3-8a), which is essential for its activity. -Amino Acids Are Found Primarily in Bacteria Although D-amino acids do not generally occur in proteins, they do serve some special functions in the structure of bacterial cell walls and peptide antibiotics. Bacterial peptidoglycans (see Fig. 6- 32) contain both D-alanine and D-glutamate. D-Amino acids arise directly from the L isomers by the action of amino acid racemases, which have pyridoxal phosphate as cofactor (see Fig. 18-6). Amino acid racemization is uniquely important to bacterial metabolism, and enzymes such as alanine racemase are prime targets for pharmaceutical agents. One such agent, L- fluoroalanine, is being tested as an antibacterial drug. Another, cycloserine, is used to treat tuberculosis. Because these inhibitors also affect some PLP-requiring human enzymes, however, they have potentially undesirable side effects. Aromatic Amino Acids Are Precursors of Many Plant Substances Phenylalanine, tyrosine, and tryptophan are converted to a variety of important compounds in plants. The rigid polymer lignin, derived from phenylalanine and tyrosine, is second only to cellulose in abundance in plant tissues. The structure of the lignin polymer is complex and not well understood. Tryptophan is also the precursor of the plant growth hormone indole-3-acetate, or auxin (Fig. 22-30a), which is important in the regulation of a wide range of biological processes in plants. FIGURE 22-30 Biosynthesis of two plant substances from amino acids. (a) Indole-3-acetate (auxin) and (b) cinnamate (cinnamon flavor). Phenylalanine and tyrosine also give rise to many commercially significant natural products, including the tannins that inhibit oxidation in wines; alkaloids such as morphine, which have potent physiological effects; and the flavoring of cinnamon oil (Fig. 22-30b), nutmeg, cloves, vanilla, cayenne pepper, and other products. Biological Amines Are Products of Amino Acid Decarboxylation Many important neurotransmitters are primary or secondary amines, derived from amino acids in simple pathways. In addition, some polyamines that form complexes with DNA are derived from the amino acid ornithine, a component of the urea cycle. A common denominator of many of these pathways is amino acid decarboxylation, another PLP-requiring reaction (see Fig. 18-6). The synthesis of some neurotransmitters is illustrated in Figure 22-31. Tyrosine gives rise to a family of catecholamines that includes dopamine, norepinephrine, and epinephrine. Levels of catecholamines are correlated with, among other things, changes in blood pressure. The neurological disorder Parkinson disease is associated with an underproduction of dopamine, and it has traditionally been treated by administering L-dopa. Overproduction of dopamine in the brain may be linked to psychological disorders such as schizophrenia. FIGURE 22-31 Biosynthesis of some neurotransmitters from amino acids. The key step is the same in each case: a PLP-dependent decarboxylation (shaded light red). Glutamate decarboxylation gives rise to γ -aminobutyrate (GABA), an inhibitory neurotransmitter. Its underproduction is associated with epileptic seizures. GABA analogs are used in the treatment of epilepsy and hypertension. Levels of GABA can also be increased by administering inhibitors of the GABA-degrading enzyme GABA aminotransferase. Another important neurotransmitter, serotonin, is derived from tryptophan in a two- step pathway. Histidine undergoes decarboxylation to histamine, a powerful vasodilator in animal tissues. Histamine is released in large amounts as part of the allergic response, and it also stimulates acid secretion in the stomach. A growing array of pharmaceutical agents are being designed to interfere with either the synthesis or the action of histamine. A prominent example is the histamine receptor antagonist cimetidine (Tagamet), a structural analog of histamine: It promotes the healing of duodenal ulcers by inhibiting secretion of gastric acid. Polyamines such as spermine and spermidine, involved in DNA packaging, are derived from methionine and ornithine by the pathway shown in Figure 22-32. The first step is decarboxylation of ornithine, a precursor of arginine (Fig. 22-12). Ornithine decarboxylase, a PLP-requiring enzyme, is the target of several powerful inhibitors used as pharmaceutical agents (see Box 6-1). FIGURE 22-32 Biosynthesis of spermidine and spermine. The PLP-dependent decarboxylation steps are shaded light red. In these reactions, S-adenosylmethionine (in its decarboxylated form) acts as a source of propylamino groups (shaded blue). Arginine Is the Precursor for Biological Synthesis of Nitric Oxide A surprise finding in the mid-1980s was the role of nitric oxide (NO) — previously known mainly as a component of smog — as an important biological messenger. This simple gaseous substance diffuses readily through membranes, although its high reactivity limits its range of diffusion to about a 1 mm radius from the site of synthesis. In humans NO plays a role in a range of physiological processes, including neurotransmission, blood clotting, and the control of blood pressure. Its mode of action is described in Box 12-2. Nitric oxide is synthesized from arginine in an NADPH- dependent reaction catalyzed by nitric oxide synthase (Fig. 22- 33), a dimeric enzyme structurally related to NADPH cytochrome P-450 reductase (see Box 21-1). The reaction is a five-electron oxidation. Each subunit of the enzyme contains one bound molecule of each of four different cofactors: FMN, FAD, tetrahydrobiopterin, and Fe3+ heme. NO is an unstable molecule and cannot be stored. Its synthesis is stimulated by interaction of nitric oxide synthase with Ca2+-calmodulin (see Fig. 12-17). FIGURE 22-33 Biosynthesis of nitric oxide. The nitrogen of the NO is derived from the guanidinium group of arginine. SUMMARY 22.3 Molecules Derived from Amino Acids Many important biomolecules are derived from amino acids. Glycine is a precursor of porphyrins. Degradation of iron-porphyrin (heme) generates bilirubin, which is converted to bile pigments with several physiological functions. Glycine and arginine give rise to creatine and phosphocreatine, an energy buffer. Glutathione, formed from three amino acids, is an important cellular reducing agent. Bacteria synthesize D-amino acids from L-amino acids in racemization reactions requiring pyridoxal phosphate. D-Amino acids are commonly found in certain bacterial walls and certain antibiotics. Plants make many substances from aromatic amino acids. The PLP-dependent decarboxylation of some amino acids yields important biological amines, including neurotransmitters and polyamines. Arginine is the precursor of nitric oxide, a biological messenger. 22.4 Biosynthesis and Degradation of Nucleotides As discussed in Chapter 8, nucleotides have a variety of important functions in all cells. They are the precursors of DNA and RNA. They are essential carriers of chemical energy — a role primarily of ATP and to some extent GTP. They are components of the cofactors NAD, FAD, S-adenosylmethionine, and coenzyme A, as well as of activated biosynthetic intermediates such as UDP- glucose and CDP-diacylglycerol. Some, such as cAMP and cGMP, are also cellular second messengers. Two types of pathways lead to nucleotides: the de novo pathways and the salvage pathways. De novo synthesis of nucleotides begins with their metabolic precursors: amino acids, ribose 5- phosphate, CO2, and NH3. Salvage pathways recycle the free bases and nucleosides released from nucleic acid breakdown. Both types of pathways are important in cellular metabolism, and both are discussed in this section. The de novo pathways for purine and pyrimidine biosynthesis seem to be nearly identical in all living organisms. Notably, the free bases guanine, adenine, thymine, cytidine, and uracil are not intermediates in these pathways; that is, the bases are not synthesized and then attached to ribose, as might be expected. The purine ring structure is built up one or a few atoms at a time, attached to ribose throughout the process. The pyrimidine ring is synthesized as orotate, attached to ribose phosphate, and then converted to the common pyrimidine nucleotides required in nucleic acid synthesis. Although the free bases are not intermediates in the de novo pathways, they are intermediates in some of the salvage pathways. Several important precursors are shared by the de novo pathways for synthesis of pyrimidines and purines. Phosphoribosyl pyrophosphate (PRPP) is important in both, and in these pathways the structure of ribose is retained in the product nucleotide, in contrast to its fate in the tryptophan and histidine biosynthetic pathways discussed earlier. An amino acid is an important precursor in each type of pathway: glycine for purines and aspartate for pyrimidines. Glutamine again is the most important source of amino groups — in five different steps in the de novo pathways. Aspartate is also used as the source of an amino group in the purine pathways, in two steps. Two other features deserve mention. First, there is evidence, especially in the de novo purine pathway, that the enzymes are present as large, multienzyme complexes or metabolons in the cell, a recurring theme in our discussion of metabolism. Second, the cellular pools of nucleotides (other than ATP) are quite small, perhaps 1% or less of the amounts required to synthesize the cell’s DNA. Therefore, cells must continue to synthesize nucleotides during nucleic acid synthesis, and in some cases, nucleotide synthesis may limit the rates of DNA replication and transcription. Because of the importance of these processes in dividing cells, agents that inhibit nucleotide synthesis have become particularly important in medicine. We examine here the biosynthetic pathways of purine and pyrimidine nucleotides and their regulation, the formation of the deoxynucleotides, and the degradation of purines and pyrimidines to uric acid and urea. We end with a discussion of chemotherapeutic agents that affect nucleotide synthesis. De Novo Purine Nucleotide Synthesis Begins with PRPP The two parent purine nucleotides of nucleic acids are adenosine 5′-monophosphate (AMP; adenylate) and guanosine 5′- monophosphate (GMP; guanylate), containing the purine bases adenine and guanine. Figure 22-34 shows the origin of the carbon and nitrogen atoms of the purine ring system, as determined by John M. Buchanan using isotopic tracer experiments in birds (who conveniently excrete excess nitrogen as insoluble uric acid, a purine analog). The detailed pathway of purine biosynthesis was worked out primarily by Buchanan and G. Robert Greenberg in the 1950s. FIGURE 22-34 Origin of the ring atoms of purines. This information was obtained from isotopic experiments with 14C- or 15N-labeled precursors. Formate is supplied in the form of N 10-formyltetrahydrofolate. In the first committed step of the pathway, an amino group donated by glutamine is attached at C-1 of PRPP (Fig. 22-35). The resulting 5-phosphoribosylamine is highly unstable, with a half- life of 30 seconds at pH 7.5. This intermediate is rapidly funneled into the next biosynthetic step, and the purine ring is subsequently built up on this structure. The pathway described here is identical in all organisms, with the exception of one step that differs in higher eukaryotes, as noted below. FIGURE 22-35 De novo synthesis of purine nucleotides: construction of the purine ring of inosinate (IMP). Each addition to the purine ring is shaded to match Figure 22-34. A er step , R symbolizes the 5-phospho- -ribosyl group on which the purine ring is built. Formation of 5-phosphoribosylamine (step ) is the first committed step in purine synthesis. Note that the product of step , AICAR, is the remnant of ATP released during histidine biosynthesis (see Fig. 22-22, step ). Abbreviations are given for most intermediates to simplify the naming of the enzymes. Step is the alternative path from AIR to CAIR occurring in higher eukaryotes. The second step is the addition of three atoms from glycine (Fig. 22-35, step ). An ATP is consumed to activate the glycine carboxyl group (in the form of an acyl phosphate) for this condensation reaction. The added glycine amino group is then formylated by N 10-formyltetrahydrofolate (step ), and a nitrogen is contributed by glutamine (step ), before dehydration and ring closure yield the five-membered imidazole ring of the purine nucleus, as 5-aminoimidazole ribonucleotide (AIR; step ). At this point, three of the six atoms needed for the second ring in the purine structure are in place. To complete the process, a carboxyl group is first added (step ). This carboxylation is unusual in that it does not require biotin, but instead uses the bicarbonate generally present in aqueous solutions. A rearrangement transfers the carboxylate from the exocyclic amino group to position 4 of the imidazole ring (step ). Steps and are found only in bacteria and fungi. In higher eukaryotes, including humans, the 5-aminoimidazole ribonucleotide product of step is carboxylated directly to carboxyaminoimidazole ribonucleotide in one step instead of two (step ). The enzyme catalyzing this reaction is AIR carboxylase. Aspartate now donates its amino group in two steps ( and ): formation of an amide bond, followed by elimination of the carbon skeleton of aspartate (as fumarate). (Recall that aspartate plays an analogous role in two steps of the urea cycle; see Fig. 18- 10.) The final carbon is contributed by N 10-formyltetrahydrofolate (step ), and a second ring closure takes place to yield the second fused ring of the purine nucleus (step ). The first intermediate with a complete purine ring is inosinate (IMP). As in the tryptophan and histidine biosynthetic pathways, the enzymes of IMP synthesis seem to be organized as large metabolons in the cell. Once again, evidence comes from the existence of single polypeptides with several functions, some catalyzing nonsequential steps in the pathway. In eukaryotic cells ranging from yeast to fruit flies to chickens, steps , , and in Figure 22-35 are catalyzed by a multifunctional protein. An additional multifunctional protein catalyzes steps and . In humans, a multifunctional enzyme combines the activities of AIR carboxylase and SAICAR synthetase (steps and ). In bacteria, these activities are found on separate proteins, but the proteins may form a metabolon. The channeling of reaction intermediates from one enzyme to the next permitted by these complexes is probably especially important for unstable intermediates such as 5-phosphoribosylamine. Conversion of inosinate to adenylate requires the insertion of an amino group derived from aspartate (Fig. 22-36); this takes place in two reactions similar to those used to introduce N-1 of the purine ring (Fig. 22-35, steps and ). A crucial difference is that GTP rather than ATP is the source of the high-energy phosphate in synthesizing adenylosuccinate. Guanylate is formed by the NAD+-requiring oxidation of inosinate at C-2, followed by addition of an amino group derived from glutamine. ATP is cleaved to AMP and PPi in the final step (Fig. 22-36). FIGURE 22-36 Biosynthesis of AMP and GMP from IMP. Purine Nucleotide Biosynthesis Is Regulated by Feedback Inhibition Four major feedback mechanisms cooperate in regulating the overall rate of de novo purine nucleotide synthesis and the relative rates of formation of the two end products, adenylate and guanylate (Fig. 22-37). The first mechanism is exerted on the first reaction that is unique to purine synthesis: transfer of an amino group to PRPP to form 5-phosphoribosylamine. This reaction is catalyzed by the allosteric enzyme glutamine-PRPP amidotransferase, which is inhibited by the end products IMP, AMP, and GMP. AMP and GMP act synergistically in this concerted inhibition. Thus, whenever either AMP or GMP accumulates to excess, the first step in its biosynthesis from PRPP is partially inhibited. FIGURE 22-37 Regulatory mechanisms in the biosynthesis of adenine and guanine nucleotides in E. coli. Regulation of these pathways differs in other organisms. In the second control mechanism, exerted at a later stage, an excess of GMP in the cell inhibits formation of xanthylate from inosinate by IMP dehydrogenase, without affecting the formation of AMP. Conversely, an accumulation of adenylate inhibits formation of adenylosuccinate by adenylosuccinate synthetase, without affecting the biosynthesis of GMP. When both products are present in sufficient quantities, IMP builds up, and it inhibits an earlier step in the pathway; this is another example of the regulatory strategy called sequential feedback inhibition. In the third mechanism, GTP is required in the conversion of IMP to AMP, whereas ATP is required for conversion of IMP to GMP (Fig. 22-36), a reciprocal arrangement that tends to balance the synthesis of the two ribonucleotides. The fourth and final control mechanism is the inhibition of PRPP synthesis by the allosteric regulation of ribose phosphate pyrophosphokinase. This enzyme is inhibited by ADP and GDP, in addition to metabolites from other pathways for which PRPP is a starting point. Pyrimidine Nucleotides Are Made from Aspartate, PRPP, and Carbamoyl Phosphate The common pyrimidine ribonucleotides are cytidine 5′- monophosphate (CMP; cytidylate) and uridine 5′-monophosphate (UMP; uridylate), which contain the pyrimidines cytosine and uracil. De novo pyrimidine nucleotide biosynthesis (Fig. 22-38) proceeds in a somewhat different manner from purine nucleotide synthesis; the six-membered pyrimidine ring is made first and then attached to ribose 5-phosphate. Required in this process is carbamoyl phosphate, also an intermediate in the urea cycle. However, in animals the carbamoyl phosphate required in urea synthesis is made in mitochondria by carbamoyl phosphate synthetase I, whereas the carbamoyl phosphate required in pyrimidine biosynthesis is made in the cytosol by a different form of the enzyme, carbamoyl phosphate synthetase II. In bacteria, a single enzyme supplies carbamoyl phosphate for the synthesis of arginine and pyrimidines. The bacterial enzyme has three separate active sites, spaced along a channel nearly 100 Å long (Fig. 22-39). Bacterial carbamoyl phosphate synthetase provides a vivid illustration of the channeling of unstable reaction intermediates between active sites so that products are formed efficiently.

FIGURE 22-38 De novo synthesis of pyrimidine nucleotides: biosynthesis of UTP and CTP via orotidylate. The pyrimidine is constructed from carbamoyl phosphate and aspartate. The ribose 5-phosphate is then added to the completed pyrimidine ring by orotate phosphoribosyltransferase. The first step in this pathway (not shown here; see Fig. 18-11a) is the synthesis of carbamoyl phosphate from CO2, NH+4, and ATP. In eukaryotes, the first step is catalyzed by carbamoyl phosphate synthetase II. FIGURE 22-39 Channeling of intermediates in bacterial carbamoyl phosphate synthetase. The reaction catalyzed by this enzyme (and its mitochondrial counterpart) is illustrated in Figure 18-11a. In this cutaway, the small and large subunits are shown in tan and blue, respectively; the tunnel between active sites (almost 100 Å long) is shown as white. In this reaction, a glutamine molecule binds to the small subunit, donating its amido nitrogen as NH+4 in a glutamine amidotransferase–type reaction. The NH+4 enters the tunnel, which takes it to a second active site, where it combines with bicarbonate in a reaction requiring ATP. The carbamate then reenters the tunnel to reach the third active site, where it is phosphorylated by ATP to carbamoyl phosphate. To solve this structure, the enzyme was crystallized with ornithine bound to the glutamine-binding site and ADP bound to the ATP-binding sites. [Data from PDB ID 1M6V, J. B. Thoden et al., J. Biol. Chem. 277:39,722, 2002.] Carbamoyl phosphate reacts with aspartate to yield N- carbamoylaspartate in the first committed step of pyrimidine biosynthesis (Fig. 22-38). This reaction is catalyzed by aspartate transcarbamoylase. In bacteria, this step is highly regulated, and bacterial aspartate transcarbamoylase is one of the most thoroughly studied allosteric enzymes (see below). By removal of water from N-carbamoylaspartate, a reaction catalyzed by dihydroorotase, the pyrimidine ring is closed to form L- dihydroorotate. This compound is oxidized to the pyrimidine derivative orotate, a reaction in which NAD+ is the ultimate electron acceptor. In eukaryotes, the first three enzymes in this pathway — carbamoyl phosphate synthetase II, aspartate transcarbamoylase, and dihydroorotase — are part of a single trifunctional protein. The protein, known by the acronym CAD, contains three identical polypeptide chains (each of Mr230,000), each with active sites for all three reactions. This suggests that metabolons may be the rule in this pathway. Once orotate is formed, the ribose 5-phosphate side chain, provided once again by PRPP, is attached to yield orotidylate (Fig. 22-38). Orotidylate is then decarboxylated to uridylate, which is phosphorylated to UTP. CTP is formed from UTP by the action of cytidylate synthetase, by way of an acyl phosphate intermediate (consuming one ATP). The nitrogen donor is normally glutamine, although the cytidylate synthetases in many species can use NH+4 directly. Pyrimidine Nucleotide Biosynthesis Is Regulated by Feedback Inhibition Regulation of the rate of pyrimidine nucleotide synthesis in bacteria occurs in large part through aspartate transcarbamoylase (ATCase), which catalyzes the first reaction in the sequence and is inhibited by CTP, the end product of the sequence (Fig. 22-38). The bacterial ATCase molecule consists of six catalytic subunits and six regulatory subunits (see Fig. 6-36). The catalytic subunits bind the substrate molecules, and the allosteric subunits bind the allosteric inhibitor, CTP. The entire ATCase molecule, as well as its subunits, exists in two conformations, active and inactive. When CTP is not bound to the regulatory subunits, the enzyme is maximally active. As CTP accumulates and binds to the regulatory subunits, they undergo a change in conformation. This change is transmitted to the catalytic subunits, which then also shi to an inactive conformation. ATP prevents the changes induced by CTP. Figure 22-40 shows the effects of the allosteric regulators on the activity of ATCase. FIGURE 22-40 Allosteric regulation of aspartate transcarbamoylase by CTP and ATP. Addition of 0.8mM CTP, the allosteric inhibitor of ATCase, increases the K0.5 for aspartate (lower curve), thereby reducing the rate of conversion of aspartate to N-carbamoylaspartate. ATP at 0.6mM fully reverses this inhibition by CTP (middle curve). Nucleoside Monophosphates Are Converted to Nucleoside Triphosphates Nucleotides to be used in biosynthesis are generally converted to nucleoside triphosphates. The conversion pathways are common to all cells. Phosphorylation of AMP to ADP is promoted by adenylate kinase, in the reaction AT P +  AM P ⇌  2ADP The ADP so formed is phosphorylated to ATP by the glycolytic enzymes or through oxidative phosphorylation. ATP also brings about the formation of other nucleoside diphosphates by the action of a class of enzymes called nucleoside monophosphate kinases. These enzymes, which are generally specific for a particular base but nonspecific for the sugar (ribose or deoxyribose), catalyze the reaction AT P +  NM P ⇌  ADP+ NDP The efficient cellular systems for rephosphorylating ADP to ATP (ATP synthase; Chapter 19) tend to remove ADP and pull this reaction in the direction of products. Nucleoside diphosphates are converted to triphosphates by the action of a ubiquitous enzyme, nucleoside diphosphate kinase, which catalyzes the reaction NT PD +  NDPA ⇌  NDPD +  NT PA This enzyme is notable in that it is not specific for the base (purines or pyrimidines) or the sugar (ribose or deoxyribose). This nonspecificity applies to both phosphate acceptor (A) and donor (D), although the donor (NT PD) is almost invariably ATP because it is present in higher concentration than other nucleoside triphosphates under aerobic conditions. Ribonucleotides Are the Precursors of Deoxyribonucleotides Deoxyribonucleotides, the building blocks of DNA, are derived from the corresponding ribonucleotides by direct reduction at the 2′-carbon atom of the D-ribose to form the 2′-deoxy derivative. For example, adenosine diphosphate (ADP) is reduced to 2′- deoxyadenosine diphosphate (dADP), and GDP is reduced to dGDP. This reaction is somewhat unusual in that the reduction occurs at a nonactivated carbon; no closely analogous chemical reactions are known. The reaction is catalyzed by ribonucleotide reductase, best characterized in E. coli, in which its substrates are ribonucleoside diphosphates. The reduction of the D-ribose portion of a ribonucleoside diphosphate to 2′-deoxy-D-ribose requires a pair of hydrogen atoms, which are ultimately donated by NADPH via an intermediate hydrogen-carrying protein, thioredoxin. This ubiquitous protein serves a similar redox function in photosynthesis (see Fig. 20-37) and other processes. Thioredoxin has pairs of — SH groups that carry hydrogen atoms from NADPH to the ribonucleoside diphosphate. Its oxidized (disulfide) form is reduced by NADPH in a reaction catalyzed by thioredoxin reductase (Fig. 22-41), and reduced thioredoxin is then used by ribonucleotide reductase to reduce the nucleoside diphosphates (NDPs) to deoxyribonucleoside diphosphates (dNDPs). A second source of reducing equivalents for ribonucleotide reductase is glutathione (GSH). Glutathione serves as the reductant for a protein closely related to thioredoxin, glutaredoxin, which then transfers the reducing power to ribonucleotide reductase. FIGURE 22-41 Reduction of ribonucleotides to deoxyribonucleotides by ribonucleotide reductase. Electrons are transmitted (red arrows) to the enzyme from NADPH via (a) glutaredoxin or (b) thioredoxin. The sulfhydryl groups in glutaredoxin reductase are contributed by two molecules of bound glutathione (GSH; GSSG indicates oxidized glutathione). Note that thioredoxin reductase is a flavoenzyme, with FAD as a prosthetic group. Ribonucleotide reductase is notable in that its reaction mechanism provides the best-characterized example of the involvement of free radicals in biochemical transformations, once thought to be rare in biological systems. The enzyme in E. coli and most eukaryotes is an α2β2 dimer, with two catalytic subunits, α2, and two radical-generation subunits, β2 (Fig. 22-42). Each catalytic subunit contains two kinds of regulatory sites, as described below. The two active sites of the enzyme are formed at the interface between the catalytic (α2) and radical-generation (β2) subunits. At each active site, an α subunit contributes two sulfhydryl groups required for activity, and the β2 subunits contribute a stable tyrosyl radical. The β2 subunits also have a binuclear iron (Fe3+) cofactor that helps generate and stabilize the T yr122 radical. The tyrosyl radical is too far from the active site to interact directly with the site, but several aromatic residues form a long-range radical-transfer pathway to the active site (Fig. 22-42c). A likely mechanism for the ribonucleotide reductase reaction is illustrated in Figure 22-43. In E. coli, the sources of the required reducing equivalents for this reaction are thioredoxin and glutaredoxin, as noted above. FIGURE 22-42 Ribonucleotide reductase. (a) A schematic diagram of the subunit structures. Each catalytic subunit (α ; also called R1) contains the two regulatory sites described in Figure 22-44 and two Cys residues central to the reaction mechanism. The radical-generation subunits (β ; also called R2) each contain a critical Tyr122 residue and binuclear iron center. (b) The likely structure of α2β2. (c) The likely path of radical formation from the initial Tyr122 in a β subunit to the active-site Cys439, which is used in the mechanism shown in Figure 22-43. Several aromatic amino acid residues participate in long-range transfer of the radical from the point of its formation at Tyr122 to the active site, where the nucleotide substrate is bound. [(a) Information from L. Thelander and P. Reichard, Annu. Rev. Biochem. 48:133, 1979. (b, c) Data from PDB ID 3UUS, N. Ando et al., Proc. Natl. Acad. Sci. USA 108:21,046, 2011.] MECHANISM FIGURE 22-43 Proposed mechanism for ribonucleotide reductase. In the enzyme of E. coli and most eukaryotes, the active thiol groups are on the α subunit. The active-site radical (— X∙) is on the β subunit and in E. coli is probably a thiyl radical of Cys439 (see Fig. 22-42). Three classes of ribonucleotide reductase have been reported. Their mechanisms (where known) generally conform to the scheme in Figure 22-43, but they differ in the identity of the group supplying the active-site radical and in the cofactors used to generate it. The E. coli enzyme (class I) requires oxygen to regenerate the tyrosyl radical if it is quenched, so this enzyme functions only in an aerobic environment. Class II enzymes, found in other microorganisms, have 5′-deoxyadenosylcobalamin (see Box 17-2) rather than a binuclear iron center. Class III enzymes have evolved to function in an anaerobic environment. E. coli contains a separate class III ribonucleotide reductase when grown anaerobically; this enzyme contains an iron-sulfur cluster (structurally distinct from the binuclear iron center of the class I enzyme) and requires NADPH and S-adenosylmethionine for activity. It uses nucleoside triphosphates rather than nucleoside diphosphates as substrates. The evolution of different classes of ribonucleotide reductase for production of DNA precursors in different environments reflects the importance of this reaction in nucleotide metabolism. Regulation of E. coli ribonucleotide reductase is unusual in that not only its activity but also its substrate specificity is regulated by the binding of effector molecules. Each α subunit has two types of regulatory sites (Fig. 22-42). One type affects overall enzyme activity and binds either ATP, which activates the enzyme, or dATP, which inactivates it. The second type alters substrate specificity in response to the effector molecule — ATP, dATP, dTTP, or dGTP — that is bound there (Fig. 22-44). When ATP or dATP is bound, reduction of UDP and CDP is favored. When dTTP or dGTP is bound, reduction of GDP or ADP, respectively, is stimulated. The scheme is designed to provide a balanced pool of precursors for DNA synthesis. ATP is also a general activator for biosynthesis and ribonucleotide reduction. The presence of dATP in small amounts increases the reduction of pyrimidine nucleotides. An oversupply of the pyrimidine dNTPs is signaled by high levels of dTTP. Abundant dTTP shis the specificity to favor reduction of GDP. High levels of dGTP, in turn, shi the specificity to ADP reduction, and high levels of dATP shut the enzyme down. These effectors are thought to induce several distinct enzyme conformations with altered specificities. FIGURE 22-44 Regulation of ribonucleotide reductase by deoxynucleoside triphosphates. The overall activity of the enzyme is affected by binding at the primary regulatory site (le ). The substrate specificity of the enzyme is affected by the nature of the effector molecule bound at the second type of regulatory site, the substrate- specificity site (right). The diagram indicates inhibition or stimulation of enzyme activity with the four different substrates. The pathway from dUDP to dTMP is described below (see Figs 22-46, 22-47). These regulatory effects are accompanied by, and presumably mediated by, large structural rearrangements in the enzyme. When the active form of the E. coli enzyme (α2β2) is inhibited by the addition of the allosteric inhibitor dATP, a ringlike α4β4 structure forms, with alternating α2 and β2 subunits (Fig. 22-45). In this altered structure, the radical-forming path from β to α is disrupted and the residues in the path are exposed to solvent, effectively preventing radical transfer and thus inhibiting the reaction. The formation of ringlike α4β4 structures is reversed when dATP levels are reduced. The yeast ribonucleotide reductase also undergoes oligomerization in the presence of dATP, forming a hexameric ring structure, α6β6. FIGURE 22-45 Oligomerization of ribonucleotide reductase induced by the allosteric inhibitor dATP. At high concentrations of dATP (50μM ), ring-shaped α4β4 structures form. In this conformation, the residues in the radical-forming path are exposed to the solvent, blocking the radical reaction and inhibiting the enzyme. The oligomerization is reversed at lower dATP concentrations. [Data from PDB ID 3UUS, N. Ando et al., Proc. Natl. Acad. Sci. USA 108:21,046, 2011.] Thymidylate Is Derived from dCDP and dUMP DNA contains thymine rather than uracil, and the de novo pathway to thymine involves only deoxyribonucleotides. The immediate precursor of thymidylate (dTMP) is dUMP. In bacteria, the pathway to dUMP begins with formation of dUTP, either by deamination of dCTP or by phosphorylation of dUDP (Fig. 22-46). The dUTP is converted to dUMP by a dUTPase. The latter reaction must be efficient to keep dUTP pools low and prevent incorporation of uridylate into DNA. FIGURE 22-46 Biosynthesis of thymidylate (dTMP). The pathways are shown beginning with the reaction catalyzed by ribonucleotide reductase. Conversion of dUMP to dTMP is catalyzed by thymidylate synthase. A one-carbon unit at the hydroxymethyl (— CH2OH) oxidation level (see Fig. 18-17) is transferred from N 5,N 10- methylenetetrahydrofolate to dUMP, then reduced to a methyl group (Fig. 22-47). The reduction occurs at the expense of oxidation of tetrahydrofolate to dihydrofolate, which is unusual in tetrahydrofolate-requiring reactions. (The mechanism of this reaction is shown in Fig. 22-52.) The dihydrofolate is reduced to tetrahydrofolate by dihydrofolate reductase — a regeneration that is essential for the many processes that require tetrahydrofolate. In plants and at least one protist, thymidylate synthase and dihydrofolate reductase reside on a single, bifunctional protein. FIGURE 22-47 Conversion of dUMP to dTMP by thymidylate synthase and dihydrofolate reductase. Serine hydroxymethyltransferase is required for regeneration of the N 5,N 10-methylene form of tetrahydrofolate. In the synthesis of dTMP, all three hydrogens of the added methyl group are derived from N 5,N 10-methylenetetrahydrofolate (light red and gray). About 10% of the human population (and up to 50% of people in impoverished communities) suffers from folic acid deficiency. When the deficiency is severe, the symptoms can include heart disease, cancer, and some types of brain dysfunction. Folic acid deficiency during pregnancy can also produce neural tube defects in infants. At least some of these symptoms arise from a reduction in thymidylate synthesis, leading to an abnormal incorporation of uracil into DNA. Uracil is recognized by DNA repair pathways (described in Chapter 25) and is cleaved from the DNA. The presence of high levels of uracil in DNA leads to strand breaks that can greatly affect the function and regulation of nuclear DNA, ultimately causing the observed effects on the heart and brain, as well as increased mutagenesis that leads to cancer. Degradation of Purines and Pyrimidines Produces Uric Acid and Urea, Respectively Purine nucleotides are degraded by a pathway in which they lose their phosphate through the action of 5′-nucleotidase (Fig. 22- 48). Adenylate yields adenosine, which is deaminated to inosine by adenosine deaminase, and inosine is hydrolyzed to hypoxanthine (its purine base) and D-ribose. Hypoxanthine is oxidized successively to xanthine and then uric acid by xanthine oxidase, a flavoenzyme with an atom of molybdenum and four iron-sulfur centers in its prosthetic group. Molecular oxygen is the electron acceptor in this complex reaction. FIGURE 22-48 Catabolism of purine nucleotides. Note that primates excrete much more nitrogen as urea via the urea cycle (Chapter 18) than as uric acid from purine degradation. Similarly, fish excrete much more nitrogen as NH+4 than as urea produced by the pathway shown here. GMP catabolism also yields uric acid as an end product. GMP is first hydrolyzed to guanosine, which is then cleaved to free guanine. Guanine undergoes hydrolytic removal of its amino group to yield xanthine, which is converted to uric acid by xanthine oxidase. Uric acid is the excreted end product of purine catabolism in primates, birds, and some other animals. A healthy adult human excretes uric acid at a rate of about 0.6 g/24 h; the excreted product arises in part from ingested purines and in part from turnover of the purine nucleotides of nucleic acids. In most mammals and many other vertebrates, uric acid is degraded to allantoin by the action of urate oxidase. In other organisms the pathway is further extended, as shown in Figure 22-48. The pathways for degradation of pyrimidines generally lead to NH+ 4 production and thus to urea synthesis. The carbons of thymine are degraded to succinyl-CoA; those of cytosine and uracil are degraded to acetyl-CoA (Fig. 22-49). FIGURE 22-49 Catabolism of pyrimidines. These simplified pathways show end products but no intermediates. Genetic aberrations in human purine metabolism have been found, some with serious consequences. For example, adenosine deaminase (ADA) deficiency leads to severe immunodeficiency disease in which T lymphocytes and B lymphocytes do not develop properly. Lack of ADA leads to a 100-fold increase in the cellular concentration of dATP, a strong inhibitor of ribonucleotide reductase (Fig. 22-44). High levels of dATP produce a general deficiency of other dNTPs in T lymphocytes. The basis for B-lymphocyte toxicity is less clear. Individuals with ADA deficiency lack an effective immune system and do not survive unless treated. Current therapies include bone marrow transplants from a matched donor to replace the hematopoietic stem cells that mature into B and T lymphocytes. However, transplant recipients oen suffer a variety of cognitive and physiological problems. Enzyme replacement therapy, requiring once- or twice-weekly intramuscular injection of active ADA, is effective, but the therapeutic benefit oen declines aer 8 to 10 years; complications may then arise, including malignancies. For many people, a permanent cure requires replacing the defective gene with a functional one in bone marrow cells. ADA deficiency was one of the first targets of human gene therapy trials (in 1990). Mixed results in early trials have given way to significant successes, and gene therapy is rapidly becoming a viable path for long-term restoration of immune function for these patients. Newer approaches based on CRISPR-mediated gene editing (see Fig. 9-21) may eventually be even more effective. Purine and Pyrimidine Bases Are Recycled by Salvage Pathways Free purine and pyrimidine bases are constantly released in cells during the metabolic degradation of nucleotides. Free purines are in large part salvaged and reused to make nucleotides, in a pathway much simpler than the de novo synthesis of purine nucleotides described earlier. One of the primary salvage pathways consists of a single reaction catalyzed by adenosine phosphoribosyltransferase, in which free adenine reacts with PRPP to yield the corresponding adenine nucleotide: Adenine +  PRPP →  AM P +  PPi Free guanine and hypoxanthine (the deamination product of adenine; Fig. 22-48) are salvaged in the same way by hypoxanthine-guanine phosphoribosyltransferase. A similar salvage pathway exists for pyrimidine bases in microorganisms, and possibly in mammals. A genetic lack of hypoxanthine-guanine phosphoribosyltransferase activity, seen almost exclusively in young boys, results in a set of symptoms called Lesch-Nyhan syndrome. Children with this genetic disorder, which becomes manifest by the age of 2 years, are sometimes poorly coordinated and have intellectual deficits. In addition, they are extremely hostile and show compulsive self-destructive tendencies: they mutilate themselves by biting off their fingers, toes, and lips. The devastating effects of Lesch-Nyhan syndrome illustrate the importance of the salvage pathways. Hypoxanthine and guanine arise constantly from the breakdown of nucleic acids. In the absence of hypoxanthine-guanine phosphoribosyltransferase, PRPP levels rise and purines are overproduced by the de novo pathway, resulting in high levels of uric acid production and goutlike damage to tissue (see below). The brain is especially dependent on the salvage pathways, and this may account for the central nervous system damage in children with Lesch-Nyhan syndrome. This syndrome is another potential target for gene therapy. Excess Uric Acid Causes Gout Long thought (erroneously) to be due to “high living,” gout is a disease of the joints caused by an elevated concentration of uric acid in the blood and tissues. The joints become inflamed, painful, and arthritic, owing to the abnormal deposition of sodium urate crystals. The kidneys are also affected, as excess uric acid is deposited in the kidney tubules. Gout occurs predominantly in males. Its precise cause is not known, but it oen involves an underexcretion of urate. A genetic deficiency of one or another enzyme of purine metabolism may also be a factor in some cases. Gout is effectively treated by a combination of nutritional and drug therapies. Patients exclude foods especially rich in nucleotides and nucleic acids, such as liver or glandular products, from the diet. Major alleviation of the symptoms is provided by the drug allopurinol (Fig. 22-50), which inhibits xanthine oxidase, the enzyme that catalyzes the conversion of purines to uric acid. Allopurinol is a substrate of xanthine oxidase, which converts allopurinol to oxypurinol (alloxanthine). Oxypurinol inactivates the reduced form of the enzyme by remaining tightly bound in its active site. When xanthine oxidase is inhibited, the excreted products of purine metabolism are xanthine and hypoxanthine, which are more water-soluble than uric acid and less likely to form crystalline deposits. Allopurinol was developed by Gertrude Elion and George Hitchings, who also developed acyclovir, used in treating people with genital and oral herpes infections, and other purine analogs used in cancer chemotherapy.

FIGURE 22-50 Allopurinol, an inhibitor of xanthine oxidase. Hypoxanthine is the normal substrate of xanthine oxidase. A slight alteration in the structure of hypoxanthine (shaded light red) yields the medically effective enzyme inhibitor allopurinol. At the active site, allopurinol is converted to oxypurinol, a strong competitive inhibitor that remains tightly bound to the reduced form of the enzyme. George Hitchings, 1905–1998, and Gertrude Elion, 1918–1999. Many Chemotherapeutic Agents Target Enzymes in Nucleotide Biosynthetic Pathways The growth of cancer cells is not controlled in the same way as cell growth in most normal tissues. Cancer cells have greater requirements for nucleotides as precursors of DNA and RNA, and consequently are generally more sensitive than normal cells to inhibitors of nucleotide biosynthesis. A number of important chemotherapeutic agents — for cancer and other diseases — act by inhibiting one or more enzymes in these pathways. Several well-studied examples can illustrate productive approaches to treatment and help us understand how these enzymes work. Important targets for pharmaceutical agents include thymidylate synthase and dihydrofolate reductase, enzymes that provide the only cellular pathway for thymine synthesis (Fig. 22-51). One inhibitor that acts on thymidylate synthase, fluorouracil, is an important chemotherapeutic agent. Fluorouracil itself is not the enzyme inhibitor. In the cell, salvage pathways convert it to the deoxynucleoside monophosphate FdUMP, which binds to and inactivates the enzyme. Inhibition by FdUMP (Fig. 22-52) is a classic example of mechanism-based enzyme inactivation. Another prominent chemotherapeutic agent, methotrexate, is an inhibitor of dihydrofolate reductase. This folate analog acts as a competitive inhibitor; the enzyme binds methotrexate with about 100 times higher affinity than dihydrofolate. Aminopterin is a related compound that acts similarly. FIGURE 22-51 Thymidylate synthesis and folate metabolism as targets of chemotherapy. (a) During thymidylate synthesis, N 5,N 10- methylenetetrahydrofolate is converted to 7,8-dihydrofolate; the N 5,N 10- methylenetetrahydrofolate is regenerated in two steps (see Fig. 22-47). This cycle is a major target of several chemotherapeutic agents. (b) Fluorouracil and methotrexate are important chemotherapeutic agents. In cells, fluorouracil is converted to FdUMP, which inhibits thymidylate synthase. Methotrexate, a structural analog of tetrahydrofolate, inhibits dihydrofolate reductase; the shaded amino and methyl groups replace a carbonyl oxygen and a proton, respectively, in folate. Another important folate analog, aminopterin, is identical to methotrexate except that it lacks the shaded methyl group. Trimethoprim, a tight-binding inhibitor of bacterial dihydrofolate reductase, was developed as an antibiotic.

MECHANISM FIGURE 22-52 Conversion of dUMP to dTMP and its inhibition by FdUMP. The normal reaction mechanism of thymidylate synthase (le ). The nucleophilic sulfhydryl group contributed by the enzyme in step and the ring atoms of dUMP taking part in the reaction are shown in red; :B denotes an amino acid side chain that acts as a base to abstract a proton a er step . The hydrogens derived from the methylene group of N 5,N 10-methylenetetrahydrofolate are shaded in gray. The 1,3 hydride shi (step ) moves a hydride ion (shaded light red) from C-6 of tetrahydrofolate to the methyl group of thymidine, oxidizing tetrahydrofolate to dihydrofolate. This hydride shi is blocked when FdUMP is the substrate (right). Steps and proceed normally, but they result in a stable complex — consisting of FdUMP linked covalently to the enzyme and to tetrahydrofolate — that inactivates the enzyme. The medical potential of inhibitors of nucleotide biosynthesis is not limited to cancer treatment. All fast-growing cells (including bacteria and protists) are potential targets. Trimethoprim, an antibiotic developed by Hitchings and Elion, binds to bacterial dihydrofolate reductase nearly 100,000 times better than to the mammalian enzyme. It is used to treat certain urinary and middle-ear bacterial infections. Parasitic protists, such as the trypanosomes that cause African sleeping sickness (African trypanosomiasis), lack pathways for de novo nucleotide biosynthesis and are particularly sensitive to agents that interfere with their ability to use salvage pathways to scavenge nucleotides from the surrounding environment. Allopurinol (Fig. 22-50) and several similar purine analogs have shown promise for the treatment of African trypanosomiasis and related afflictions. See Box 6-1 for another approach to combating African trypanosomiasis, made possible by advances in our understanding of metabolism and enzyme mechanisms. SUMMARY 22.4 Biosynthesis and Degradation of Nucleotides The purine ring system is built up step by step, beginning with 5-phosphoribosylamine. The amino acids glutamine, glycine, and aspartate furnish all the nitrogen atoms of purines. Two ring- closure steps form the purine nucleus. Purine biosynthesis is regulated by an elaborate system of feedback inhibition. Pyrimidines are synthesized from carbamoyl phosphate and aspartate, and ribose 5-phosphate is then attached to yield the pyrimidine ribonucleotides. Pyrimidine biosynthesis is regulated by feedback inhibition of aspartate transcarbamoylase. Nucleoside monophosphates are converted to their triphosphates by enzymatic phosphorylation reactions. Ribonucleotides are converted to deoxyribonucleotides by ribonucleotide reductase, an enzyme with novel mechanistic and regulatory characteristics. The thymine nucleotides are derived from dCDP and dUMP. Uric acid and urea are the end products of purine and pyrimidine degradation. Free purines can be salvaged and rebuilt into nucleotides. Genetic deficiencies in certain salvage enzymes cause serious disorders such as Lesch-Nyhan syndrome. Accumulation of uric acid crystals in the joints, possibly caused by another genetic deficiency, results in gout. Enzymes of the nucleotide biosynthetic pathways are targets for an array of chemotherapeutic agents used to treat cancer and other diseases. Chapter Review KEY TERMS Terms in bold are defined in the glossary. nitrogen fixation nitrification denitrification anammox symbionts nitrogenase complex FeMo cofactor ferredoxin leghemoglobin glutamate glutamine glutamine synthetase glutamate synthase glutamine amidotransferases 5-phosphoribosyl-1-pyrophosphate (PRPP) porphyrin porphyria bilirubin phosphocreatine creatine glutathione (GSH) auxin dopamine norepinephrine epinephrine γ -aminobutyrate (GABA) serotonin histamine spermine spermidine ornithine decarboxylase de novo pathway salvage pathway inosinate (IMP) carbamoyl phosphate synthetase II aspartate transcarbamoylase nucleoside monophosphate kinase nucleoside diphosphate kinase ribonucleotide reductase thioredoxin thymidylate synthase dihydrofolate reductase adenosine deaminase (ADA) deficiency Lesch-Nyhan syndrome allopurinol fluorouracil methotrexate aminopterin trimethoprim PROBLEMS 1. ATP Consumption by Root Nodules in Legumes Bacteria residing in the root nodules of the pea plant consume more than 20% of the ATP produced by the plant. Suggest why these bacteria consume so much ATP. 2. Nitrate Fertilizers and Oceanic Dead Zones Farmers apply industrially fixed nitrogen, in the form of ammonia or nitrate, to agricultural fields worldwide to increase crop yields. Agricultural runoff feeds into rivers and creates large hypoxic dead zones at the point where rivers meet oceans. How does an increase in soluble fixed nitrogen create dead zones? 3. PLP Reaction Mechanisms Pyridoxal phosphate (PLP) can help catalyze transformations one or two carbons removed from the α carbon of an amino acid. The enzyme threonine synthase promotes the PLP-dependent conversion of phosphohomoserine to threonine. Suggest a mechanism for this reaction. 4. Transformation of Aspartate to Asparagine There are two routes for transforming aspartate to asparagine at the expense of ATP. Many bacteria have an asparagine synthetase that uses ammonium ion as the nitrogen donor. Mammals have an asparagine synthetase that uses glutamine as the nitrogen donor. Given that the latter requires an extra ATP (for the synthesis of glutamine), why do mammals use this route? 5. Equation for the Synthesis of Aspartate from Glucose Write the net equation for the synthesis of aspartate (a nonessential amino acid) from glucose, carbon dioxide, and ammonia. 6. Asparagine Synthetase Inhibitors in Leukemia Therapy Mammalian asparagine synthetase is a glutamine- dependent amidotransferase. Efforts to identify an effective inhibitor of human asparagine synthetase for use in chemotherapy for patients with leukemia have focused not on the amino-terminal glutaminase domain but on the carboxyl-terminal synthetase active site. Explain why the glutaminase domain is not a promising target for a useful drug. 7. Phenylalanine Hydroxylase Deficiency and Diet Tyrosine is normally a nonessential amino acid, but individuals with a genetic defect in phenylalanine hydroxylase require tyrosine in their diet for normal growth. Explain. 8. Arginine Biosynthesis The first step of arginine biosynthesis from glutamate acetylates glutamate on the α - amino group. A subsequent step late in the same pathway removes the added acetyl group. What chemical problem is solved by adding and then removing an acetyl group, with none of the acetyl atoms appearing in the arginine product of the pathway? 9. Cofactors for One-Carbon Transfer Reactions Most one- carbon transfers are promoted by one of three cofactors: biotin, tetrahydrofolate, or S-adenosylmethionine. S- Adenosylmethionine generally serves as a methyl group donor; the transfer potential of the methyl group in N5- methyltetrahydrofolate is insufficient for most biosynthetic reactions. However, one example of the use of N5- methyltetrahydrofolate in methyl group transfer is in methionine formation by the methionine synthase reaction; methionine is the immediate precursor of S- adenosylmethionine (see Fig. 18-18). Explain how the methyl group of S-adenosylmethionine can be derived from N5-methyltetrahydrofolate, even though the transfer potential of the methyl group in N5- methyltetrahydrofolate is one-thousandth of that of S- adenosylmethionine. 10. Concerted Regulation in Amino Acid Biosynthesis Various products of glutamine metabolism independently modulate the glutamine synthetase of E. coli (see Fig. 22-8). In this concerted inhibition, the extent of enzyme inhibition is greater than the sum of the separate inhibitions caused by each product. For E. coli grown in a medium rich in histidine, what is the advantage of concerted inhibition? 11. Relationship between Folic Acid Deficiency and Anemia Folic acid deficiency, believed to be the most common vitamin deficiency, causes a type of anemia in which hemoglobin synthesis is impaired and erythrocytes do not mature properly. What is the metabolic relationship between hemoglobin synthesis and folic acid deficiency? 12. Synthesis of Polyamines The metabolic amino acid ornithine is a direct precursor of the polyamine putrescine, shown here. H3 + N— CH2— CH2— CH2— CH2— + NH3 Subsequent reactions convert putrescine to spermine and spermidine. What type of reaction is required to convert ornithine to putrescine, and what enzymatic cofactor is needed? 13. Nucleotide Biosynthesis in Amino Acid Auxotrophic Bacteria Wild-type E. coli cells can synthesize all 20 common amino acids, but some mutants, called amino acid auxotrophs, are unable to synthesize a specific amino acid and require its addition to the culture medium for optimal growth. Besides their role in protein synthesis, some amino acids are also precursors for other nitrogenous cell products. Consider the three amino acid auxotrophs that are unable to synthesize glycine, glutamine, and aspartate, respectively. For each mutant, what nitrogenous products other than proteins would the cell fail to synthesize? 14. Inhibitors of Nucleotide Biosynthesis Suggest a mechanism for the inhibition of alanine racemase by L- fluoroalanine. 15. Mode of Action of Sulfa Drugs Some bacteria require p-aminobenzoate in the culture medium for normal growth, and their growth is severely inhibited by the addition of sulfanilamide, one of the earliest sulfa drugs. Moreover, in the presence of this drug, 5-aminoimidazole-4-carboxamide ribonucleotide (AICAR; see Fig. 22-35) accumulates in the culture medium. Addition of excess p-aminobenzoate reverses these effects. a. What is the role of p-aminobenzoate in these bacteria? (Hint: See Fig. 18-16.) b. Why does AICAR accumulate in the presence of sulfanilamide? c. Why does addition of excess p-aminobenzoate reverse the inhibition and accumulation? 16. Purine Biosynthesis Which atoms of the purine ring derive from the amide nitrogen of glutamine? a. N-1 b. N-3 c. N-7 d. N-9 17. Pathway of Carbon in Pyrimidine Biosynthesis Predict the locations of 14C in orotate isolated from cells grown on a small amount of uniformly labeled [14C]succinate. Justify your prediction. 18. Nucleotides as Poor Sources of Energy Under starvation conditions, organisms can use proteins and amino acids as sources of energy. Deamination of amino acids produces carbon skeletons that can enter the glycolytic pathway and the citric acid cycle to produce energy in the form of ATP. Nucleotides are not similarly degraded for use as energy- yielding fuels. What observations about cellular physiology support this statement? What aspect of the structure of nucleotides makes them a relatively poor source of energy? 19. Treatment of Gout Physicians use allopurinol (see Fig. 22-50), an inhibitor of xanthine oxidase, to treat chronic gout. Explain the biochemical basis for this treatment. Patients treated with allopurinol sometimes develop xanthine stones in the kidneys, although the incidence of kidney damage is much lower than in untreated gout. Explain this observation in light of these solubilities in urine: uric acid, 0.15 g/L; xanthine, 0.05 g/L; and hypoxanthine, 1.4 g/L. 20. Antibiotics That Inhibit Dihydrofolate Reductase Trimethoprim, a commonly used antibiotic, inhibits the bacterial form of dihydrofolate reductase much more than it inhibits the mammalian enzyme. What metabolic processes described in this chapter are affected by depleting tetrahydrofolate? DATA ANALYSIS PROBLEM 21. Use of Modern Molecular Techniques to Determine the Synthetic Pathway of a Novel Amino Acid Most of the biosynthetic pathways described in this chapter were determined before the development of recombinant DNA technology and genomics, so the techniques were quite different from those that researchers would use today. Here we explore an example of the use of modern molecular techniques to investigate the pathway of synthesis of a novel amino acid, (2S)-4-amino-2-hydroxybutyrate (AHBA). The techniques mentioned here are described in various places in the text; this problem is designed to show how they can be integrated in a comprehensive study. AHBA is a γ -amino acid that is a component of some aminoglycoside antibiotics, including the antibiotic butirosin. Antibiotics modified by the addition of an AHBA residue are oen more resistant to inactivation by bacterial antibiotic-resistance enzymes. As a result, understanding how AHBA is synthesized and added to antibiotics is useful in the design of pharmaceuticals. In an article published in 2005, Li and coworkers describe how they determined the synthetic pathway of AHBA from glutamate. a. Briefly describe the chemical transformations needed to convert glutamate to AHBA. At this point, don’t be concerned about the order of the reactions. Li and colleagues began by cloning the butirosin biosynthetic gene cluster from the bacterium Bacillus circulans, which makes large quantities of butirosin. They identified five genes that are essential for the pathway: btrI, btrJ, btrK, btrO, and btrV. They cloned these genes into E. coli plasmids that allow overexpression of the genes, producing proteins with “histidine tags” fused to their amino termini to facilitate purification (see p. 313). The predicted amino acid sequence of the BtrI protein showed strong homology to known acyl carrier proteins (see Fig. 21-5). Using mass spectrometry, Li and colleagues found a molecular mass of 11,812 for the purified BtrI protein (including the His tag). When the purified BtrI was incubated with coenzyme A and an enzyme known to attach CoA to other acyl carrier proteins, the majority molecular species had an Mr of 12,153. b. How would you use these data to argue that BtrI can function as an acyl carrier protein with a CoA prosthetic group? Using standard terminology, Li and coauthors called the form of the protein lacking CoA apo-BtrI and the form with CoA (linked as in Fig. 21-5) holo-BtrI. When holo-BtrI was incubated with glutamine, ATP, and purified BtrJ protein, the holo-BtrI species of Mr12,153 was replaced with a species of Mr12,281, corresponding to the thioester of glutamate and holo- BtrI. Based on these data, the authors proposed the following structure for the Mr12,281 species, γ - glutamyl-S-BtrI: c. What other structure(s) is (are) consistent with the data above? d. Li and coauthors argued that the structure shown here (γ -glutamyl-S-BtrI) is likely to be correct because the α -carboxyl group must be removed at some point in the synthetic process. Explain the chemical basis of this argument. (Hint: See Fig. 18-6, reaction C.) The BtrK protein showed significant homology to PLP- dependent amino acid decarboxylases, and BtrK isolated from E. coli was found to contain tightly bound PLP. When γ -glutamyl-S-BtrI was incubated with purified BtrK, a molecular species of Mr12,240 was produced. e. What is the most likely structure of this species? f. When the investigators incubated glutamate and ATP with purified BtrI, BtrJ, and BtrK, they found a molecular species of Mr12,370. What is the most likely structure of this species? Hint: Remember that BtrJ can use ATP to γ -glutamylate nucleophilic groups. Li and colleagues found that BtrO is homologous to monooxygenase enzymes (see Box 21-1) that hydroxylate alkanes, using FMN as a cofactor, and BtrV is homologous to an NAD(P)H oxidoreductase. Two other genes in the cluster, btrG and btrH, probably encode enzymes that remove the γ - glutamyl group and attach AHBA to the target antibiotic molecule. g. Based on these data, propose a plausible pathway for the synthesis of AHBA and its addition to the target antibiotic. Include the enzymes that catalyze each step and any other substrates or cofactors needed (ATP, NAD, etc.). References Li, Y., N.M. Llewellyn, R. Giri, F. Huang, and J.B. Spencer. 2005. Biosynthesis of the unique amino acid side chain of butirosin: possible protective-group chemistry in an acyl carrier protein– mediated pathway. Chem. Biol. 12:665–675.