*Corresponding author. Mailing address: Department of Biology, University of Copenhagen, Ole Maaløes Vej 5, DK-2200 Copenhagen N, Denmark. E-mail:
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Nucleotides are essential components of life. They constitute the building blocks of nucleic acids, RNA and DNA, and serve as cofactors in sugar and lipid metabolism, in polyamine biosynthesis, in methylation reactions, and as carriers of energy. They are also constituents of the redox coenzymes, NAD(P) and FAD, and of signal molecules such as (p)ppGpp, which is involved in coordinating RNA synthesis with protein synthesis (see Table 1). The nucleotides are either ribo- or deoxyribonucleotides, and they are found either as mono-, di-, or triphosphates. The triphosphates are substrates for RNA and DNA polymerases. The mono- and diphosphates are intermediates in the biosynthetic reactions and they also arise from hydrolytic and/or phosphorolytic breakdown of nucleic acids or other metabolic activities either inside the cells or outside by decay of biological material (Fig. 1). Nucleobases and nucleosides are not intermediates in the pathways of purine and pyrimidine biosynthesis but, if they become available as breakdown products of nucleotides, they may be utilized, in which cases they enter the nucleotide pools via the so-called salvage pathways, which are sequences of enzymatic reactions, including the transport functions (Fig. 1).
Deoxyribonucleotides are made exclusively by reduction of the corresponding ribonucleotides, either at the diphosphate- or the triphosphate level, depending on the growth condition. However, the uracil ring of dUMP must be methylated at the 5-position to form dTMP, which is specific for the deoxyribonucleotide pool and DNA. Escherichia coli and Salmonella enterica do not contain deoxyribonucleoside kinases, except for thymidine kinase. Instead, in salvage reactions the pentose and nucleobase moieties of deoxynucleosides are generally first split from each other by phosphorolytic cleavage of the N-glycosyl bond, then the pentose sugar is catabolized and the nucleobase salvaged by a PRTase. Thymine and cytosine constitute special cases: no known PRTase utilizes either thymine or cytosine as a substrate. Therefore, cytosine can reenter the nucleotide pool following deamination to uracil and thymine only by the reverse reaction of thymidine phosphorylase, which converts thymine and deoxyribose 1-phosphate to thymidine, which is subsequently phosphorylated by thymidine kinase. Similar reverse phosphorolytic reactions followed by ribonucleoside kinase reactions could in principle form the other ribonucleosides, but they do not occur under normal growth conditions due to low intracellular concentrations of ribose 1-phosphate. The pathways of nucleotide metabolism include reactions that interconvert purine and pyrimidine nucleotides, nucleosides, and nucleobases. Generally, auxotrophic mutants blocked by single lesions in the pathways for de novo synthesis of either purine- or pyrimidine nucleotides can survive if but a single purine or pyrimidine compound is present in the growth medium.
Nucleotide biosynthesis is tightly controlled by feedback loops and by considerable cross talk between steps in the pathways for pyrimidine and purine nucleotide biosynthesis. Thus, purine ribonucleotides are involved in regulation of the pyrimidine nucleotide biosynthetic enzymes both at the level of gene expression and as substrates or effectors of the enzymes. There is also a strong link to the macromolecular biosynthesis machinery, because expression of the pyrimidine biosynthetic genes in E. coli and Salmonella is regulated primarily by changes in the transcription kinetics of RNA polymerase and interplay between the translating ribosomes and the pools of nucleoside triphosphates. Moreover, RNA breakdown and other metabolic activities that generate CMP or CDP supply the majority of the ribonucleoside diphosphates used for the formation of pyrimidine deoxyribonucleotides and, hence, for DNA synthesis under aerobic growth conditions.
Nucleosides and nucleobases with exocyclic amino groups (e.g., adenosine, cytidine, and cytosine) are excellent sources of nitrogen and are rapidly deaminated in the cells. The purine ring is generally retained, but the pyrimidine ring may be broken down to release the nitrogen content by some, but not all of enteric bacteria. The genes specifying enzymes for the degradative pathways are under nitrogen control and primarily expressed when the cells lack more readily utilizable sources of nitrogen.
In the following sections we will review the literature on the different aspects of nucleotide metabolism in enteric bacteria beginning with the synthesis of PRPP and an overview of the enzymatic reactions catalyzed by the PRTases that form the N-glycosyl bond of all nucleotides, as well as the kinases that convert the nucleoside monophosphates to di- and triphosphates. These reactions are common for salvage and de novo synthesis of nucleotides. In subsequent sections we will discuss the more specific aspects, such as the de novo biosynthetic pathways, the formation of deoxyribonucleotides, as well as nucleoside and nucleobase transport and degradation. For more detailed discussions of some aspects we refer to the chapters on nucleotide biosynthesis and metabolism in the print predecessors of EcoSal (439, 441, 681).
PRPP synthase.
5-Phosphoribosyl 1-α-diphosphate (PRPP) is the ultimate donor of the 5'-phosphoribosyl moiety of all nucleotides and participates also in histidine, tryptophan, and nicotinamide coenzyme synthesis (Fig. 4). The compound was identified by Lieberman et al. (359) in a search for a compound and an enzyme that could mediate the conversion of orotate to UMP. The enzyme, PRPP synthase, catalyzes the Mg2+-dependent reaction (see Fig. 2). PRPP synthase is encoded by the prs (or prsA) gene in E. coli and Salmonella enterica serovar Typhimurium (244, 249, 286). Despite the multiple pathways to which PRPP delivers carbon atoms (Fig. 4), PRPP-less mutants (Δprs) of E. coli have been isolated. The viability of these mutants depends on the presence of mutations in genes for nucleoside phosphorylases as well as an "upmutation" in the gene encoding guanosine kinase (gsk) in order to improve nucleoside salvage at the expense of nucleobase salvage (245). A temperature-sensitive mutation as well as a mutation with essentially the entire prs gene replaced by a kanamycin resistance-encoding cassette has been isolated (65, 250, 488). Mutants of S. enterica with a defective prs gene have also been isolated and characterized (286, 487). Increased levels of PRPP synthase have been observed during pyrimidine starvation (651). The regulatory mechanism is unknown, but the effector seems to be a uridine nucleotide (465) different from UMP (271). In addition the purine repressor, PurR with corepressors binds to the promoter region of the prs gene and transcription is increased two- to threefold in a purR mutant (216).
PRPP synthase from enteric bacteria belongs to the group of "classical" class I PRPP synthases, which are characterized by (i) the restricted use of ATP (or, in some cases, dATP) as diphosphoryl donor, (ii) allosteric regulation by (one or more) ribonucleoside diphosphates, and (iii) requirement of phosphate ion for activity. Members of the newly discovered class II PRPP synthase may utilize GTP, CTP, or UTP as diphosphoryl donors (in addition to ATP and dATP), they appear to lack allosteric regulation, and their activity is independent of phosphate ion (325, 326). PRPP synthases from E. coli and S. enterica are essentially identical, as their amino acid sequences differ by only two conservative replacements. Consequently, analyses of the kinetic mechanism of the two enzymes, which have been characterized in great detail, revealed no significant differences (173, 248, 570, 571, 573, 655, 656). The reaction proceeds by transfer of the β,γ-diphosphoryl of ATP to the C-1 hydroxyl of ribose 5-phosphate (Fig. 2). The enzyme binds one Mg2+ in the active site in addition to that complexed with ATP (570, 655). The binding of Mg2+, MgATP, and ribose 5-phosphate to the enzyme occurs in that order (570, 655). The enzyme is allosterically activated by phosphate and inhibited by ADP, which binds to both the allosteric site in competition with phosphate and to the active site in competition with ATP (173, 571, 656). The presence of ADP also causes a pronounced substrate inhibition by ribose 5-phosphate (573), which may be explained by a change in the order of binding the inhibitor to either the free enzyme or to the enzyme complexed with substrates (656).
The three-dimensional structure of the enzyme from enteric bacteria is not known but is expected to be very similar to the crystal structure of the enzyme from Bacillus subtilis (146). That enzyme is a hexamer, composed as a trimer of dimers. The subunit has a two-domain structure related by a twofold symmetry. The peptide chain fold of these domains is similar to the PRPP binding domain common to all type I PRTases (see below). This is in agreement with the finding that PRPP synthase in the C-terminal domain harbors the sequence motif corresponding to a PRPP, or ribose 5-phosphate binding site homologous to that of the type I PRTases (248, 552). The active site of PRPP synthase is situated at the interface between the two domains of the subunit. The allosteric site, however, is at the dimer interface and binds both phosphate and ADP in competition. The two metal-ion sites for binding of Mg2+ and MgATP have also been located in the active site of the crystallized B. subtilis PRPP synthase (147). Mutational analysis of the aspartate-rich PRPP binding motif of the E. coli enzyme indicated that a metal ion is involved in binding of ribose 5-phosphate to the enzyme and that the acidic residues of the sequence motif play a role in both ribose 5-phosphate binding and metal-ion specificity (660).
Ribose bisphosphokinase.
A second enzyme capable of synthesizing PRPP has been identified. This enzyme, ribose bisphosphokinase (EC 2.7.4.23), catalyzes the following reaction: ribose 1,5-bisphosphate + ATP → PRPP + ADP (251). The enzyme is encoded by the phnN gene, which belongs to the Pho regulon. The relevance of this enzyme to overall PRPP metabolism remains to be established, but it is not able to support growth of cells deleted for the prs gene (246) and the metabolic origin of the substrate ribose 1,5-bisphosphate is unknown. Furthermore, most serovars of S. enterica do not have the enzyme.
Phosphoribosyltransferases.
The literature on structure and function of PRTases has been reviewed several times (193, 531, 549). In short, although they catalyze similar reactions, the different PRTases show little sequence similarity aside from a short region of 12 residues, the PRPP binding site (or the PRPP motif) also found in PRPP synthase, which—except in UPRTase—consists of two acidic residues (DD or ED) flanked by 5 hydrophobic residues on either side (225, 248), and characteristic of each type of reaction (373). However, all PRTases contain a common structural core, which is built as a sheet of five parallel β-strands connected by loops and helices that carries the active site. The PRPP motif is situated in a loop (the PRPP-loop) that connects β3 and α3, and this loop embraces the 5-phosphoribosyl moieties of PRPP as well as the nucleotide products. The active site is covered by a hood structure that is involved in binding or supplying the cosubstrate, and a flexible loop that contributes catalytic residues and closes the site during catalysis to prevent access of water to the reactive intermediate or transition state. The enzymes are dimers or tetramers in their functional state and the flexible loop may fold into the active sites of the same subunit or a neighboring subunit. In addition, a non-proline cis-peptide bond, found in the so-called PPi loop, which connects β1 and α2 and holds the β-phosphate of PRPP (221), may undergo obligatory cis-trans isomerization during catalysis (222, 223). The hood subdomain may be formed from the C-terminal or N-terminal or both and may be small, as in OPRTase (526), or large, as in glutamine PRPP amidotransferase, in which the hood constitutes the glutaminase part of the enzyme (553). All of the enzymes probably catalyze the reaction by a sequential, dissociative SN1-like mechanism in which the C1–OPP bond of PRPP essentially is broken before the C1–N bond to incoming nucleophile is beginning to form and the phosphoribosyl moiety adopting an oxycarbenium-like transition state with the positive charge distributed between C-1 and O-4 of the ribose ring (531, 574).
Nucleoside phosphorylases.
These reactions are readily reversible and thermodynamics favor nucleoside formation over nucleoside cleavage, but nucleoside formation does not prevail in vivo because of the shortage of (deoxy)ribose 1-phosphate unless another (deoxy)nucleoside is present in the growth medium and phosphorolytically cleaved by the proper nucleoside phosphorylase (234, 280). The formation of the deoxyribonucleoside, thymidine from thymine and deoxyribose 1-phosphate in thymine-requiring (thyA) mutants, occurs because thymine-requiring mutants produce and accumulate deoxyribose 1-phosphate at high concentration because of the vigorous activity of ribonucleotide reductase at low pools of dTTP, but such synthesis of thymidine does not take place in wild-type bacteria (280).
The nucleoside monophosphates are phosphorylated to the diphosphate level by specific monophosphate kinases. The diphosphates are further converted to the triphosphate level by a nonspecific nucleoside diphosphate kinase or by the substrate level phosphorylation catalyzed by pyruvate kinase, or, in the case of ADP and aerobic growth conditions, by oxidative phosphorylation.
Nucleoside monophosphate kinases.
The nucleoside monophosphates are all phosphorylated by specific kinases. The γ-phosphoryl of the nucleoside triphosphate donor is transferred to the acceptor nucleoside monophosphate by the general scheme:
in which N is the nucleobase specific for the kinase. Only conditional mutations, temperature sensitive or cold sensitive, are described for the structural genes encoding AMP kinase (adk) (74, 176), GMP kinase (gmk) (165), UMP kinase (pyrH) (264), and dTMP kinase (tmk) (128). Therefore, these enzymes seem essential for growth and the monophosphate mandatory intermediates in the synthesis of the corresponding nucleoside di- and triphosphates. Mutants deleted for the cmk gene encoding CMP kinase are viable, because CMP is not an intermediate in the de novo pathway for the synthesis of CTP. However, the CMP kinase seems necessary for rescuing CMP that arises from intracellular, hydrolytic RNA breakdown and lipopolysaccharide and phospholipid metabolism. The cold-sensitive phenotype of CMP kinase mutations (37) and the reduced dTTP and dCTP pools, resulting in a slow DNA chain elongation kinetics (158), may be ascribed to the impaired ability of cmk mutants to convert CMP to CDP. Adenylate kinase is closely linked to the "energy charge," the index that describes the phosphorylation state of adenyl nucleotides. In principle, a similar energy charge exists for all nucleotides, but the classic understanding of this term relates to adenylate kinase and its substrates (97, 178).
The monophosphate kinases are all highly specific for the nucleobase of the phosphate acceptor substrate. UMP kinase is also specific for the ribosyl moiety (541), but AMP kinase (520), GMP kinase (462), and CMP kinase (78) also act on the corresponding deoxyribonucleoside monophosphates. The kinase that works only on deoxyribonucleoside monophosphates in E. coli and S. enterica is dTMP kinase, which phosphorylates both dTMP and dUMP (434). AMP kinase (46, 47, 419, 420), GMP kinase (227), and CMP kinase (72) are closely related in structure and function. The monomer is 200 to 230 amino acid residues long with a molecular weight of about 25 kDa. These kinases share a common structural fold formed by a core domain, comprising a central β-sheet with the strands connected by α-helices, a so-called LID domain that closes over the active site and an NMP binding domain, which is responsible for the specific binding of the nucleoside monophosphate. Depending on the presence or absence of inserts in the LID region or the NMP binding domain, the kinases are separated into two families: short and long kinases (159). AMP kinase and CMP kinase are both classified as long kinases and their characteristic inserts are in the LID region (159) and the NMP binding region (72), respectively. The oligomeric arrangement of subunits varies. AMP kinase and CMP kinase are both monomeric enzymes, whereas GMP kinase in solution has a higher oligomerization state (462) and is probably active as a dimer (227).
The phosphate donor is generally considered to be ATP, but the specificity toward this substrate is less strict than the specificity toward the nucleoside monophosphate. Structures exist of the nucleoside monophosphate kinases in complex with their substrates and analogues of the phosphoryl donor as well as the bisubstrate analogues Ap5A, Ap5G, and Ap5dT in the case of AMP kinase (419), GMP kinase (226). and dTMP kinase (338), respectively. Binding of a substrate molecule to the NMP site results in conformational changes in the NMP binding domain, which serves to provide close contacts between enzyme residues and the substrate molecule and confers specificity toward the nucleobase. Binding of the nucleoside triphosphate results in closing of the LID over the central nucleotide binding cleft and the active site region that catalyzes the phosphotransfer reaction (47, 78, 226). The catalytic residues, which interact with the phosphates of the substrates, are conserved to a large extent between the three kinds of kinases. A region in the N-terminal of the polypeptides, designated the P-loop, carries a short Walker type A sequence motif that is preceded by a β-strand and followed by an α-helix. This conserved structure is found with slight variations in the sequence motif in other kinases with a wide range of phosphoryl acceptor substrates and is known as the classical mononucleotide binding fold (596). The P-loop harbors the central catalytic lysine and serine/threonine residues that act to guide the γ-phosphoryl group during the phosphotransfer reaction (86).
UMP kinase differs from the above group of nucleoside monophosphate kinases both with respect to structure and mechanism. This enzyme is a homohexamer of polypeptides, which show structural resemblance to the amino acid kinases (71), and it does not have a Walker type A motif in the amino acid sequence to suggest the existence of a P-loop similar to the other nucleoside monophosphate kinases. Furthermore, in contrast to the other nucleoside monophosphate kinases, the activity of UMP kinase is regulated by allosteric mechanisms (148) as discussed in more detail under the section dealing with pyrimidine nucleotide synthesis.
Nucleoside diphosphate kinase.
The level of nucleoside diphosphate kinase is low during anaerobic growth, and the activity is taken over by pyruvate kinase (518). Other enzymes as well have been shown to catalyze nucleoside diphosphate kinase-like reactions. Among these are: polyphosphate kinase (330), AMP kinase (45, 371, 657), succinyl coenzyme A synthase (426), and amino acyl-tRNA synthetases (339). Mutation or deletion of ndk results in a mutator phenotype, presumably caused by imbalance between the deoxyribonucleotide pools (237, 371, 542, 684) even under anaerobiosis (542), indicates that these other enzymes cannot fully replace the nucleoside diphosphate kinase in maintaining optimal deoxyribonucleotide pools. However, E. coli strains defective in ndk and the two genes encoding the pyruvate kinase isozymes A and B, pykA and pykF, or scs, encoding succinyl coenzyme A synthase, are still able to grow even though the nucleotide pools are altered (372). The nucleoside diphosphate kinase of E. coli is a homotetramer, a dimer of dimers, but different oligomeric states of the enzyme, all composed by the same dimeric unit of monomers, prevail in other organisms (415). The reaction follows Ping-Pong kinetics with a phosphohistidine intermediate (5, 337). The structure of nucleoside diphosphate kinase in complex with ADP and beryllium- or aluminum fluoride (676) showed that a hydrogen bond between the 3'-hydroxyl of the ribosyl and the oxygen of the β,γ-phosphoester bond is formed within the substrate during both binding and catalysis. This hydrogen bond is proposed to explain the lack of activity of nucleoside diphosphate kinase with a number of nucleotide analogues lacking a 3'-hydroxyl group (675).
Nucleoside diphosphate kinase has also been ascribed different roles in bacteriophage T4 infection (542) and removal of uracil from DNA, but the precise functions of the enzyme/protein in these activities are not fully understood (41, 182, 329, 489).
Synthesis of the purine ring.
Enzymes of IMP de novo synthesis.
IMP cyclohydrolase catalyzes the final step in the purine de novo pathway, the dehydration of FAICAR to generate IMP. Little is known of enteric AICAR transformylase-IMP cyclohydrolase, but the enzyme from bacteria and eukaryotic organisms appear to be homologous, i.e., they are all bifunctional. Studies of human and avian AICAR transformylase-IMP cyclohydrolase have revealed a C-terminal domain containing AICAR transformylase activity and an NH2 domain containing IMP cyclohydrolase activity (189, 665). A similar domain structure has been shown for the enzyme from the thermophile Thermotoga maritima (24). Interpretation of results of complementation studies with mutants of E. coli has indicated an identical domain arrangement (3).
Repression is achieved by binding of PurR to an operator site, the Pur box, which is a 16-bp palindromic sequence. Its consensus sequence is ACGCAAACGTTTGCGT. This sequence, in nearly conserved form, is located at various distances in front of the transcription start sites of all of the operons specifying the purine de novo enzymes (cvpA purF, purHD, purMN, purT, purL, purEK, purC, and purB) as well as purA, guaBA, and purR itself. The position of the central nucleotide varies between position −90 and 232 relative to the transcription initiation nucleotide. The purA and purR operons have two operator sites. The repression exceeds 10-fold for the genes specifying IMP synthesis, whereas repression of purB is approximately 3-fold and purR transcription is regulated two- to threefold (397, 509, 510). The corepressors are hypoxanthine and guanine, i.e., 6-oxopurines. Thus, addition or intracellular accumulation of these compounds causes repression of the synthesis of the first four enzymes of the de novo pathway of S. enterica (243). PurR is a dimer of 38-kDa subunits (341 amino acids). Three-dimensional structures of several PurR forms have been solved, including a ternary complex consisting of PurR-hypoxanthine-purF operator, PurR-guanine-purF operator, and the ligand-free PurR corepressor domain (534, 535). The subunit consists of an NH2-terminal domain of 60 residues with DNA binding capability and a C-terminal domain to which the corepressors bind with KD values of 1.5 μM (guanine) or 9.3 μM (hypoxanthine), and which is also responsible for dimerization. The C-terminal domain itself consists of NH2- and C-terminal subdomains. The DNA binding domain contains a helix-turn-helix domain as well as a hinge-helix domain. Binding of a corepressor results in large conformational changes of the corepressor binding domain, resulting in juxtaposition of the dimer-hinge helix domains, which interdigitate the minor grove and cause a 45° or more kinking of the DNA at the central CpG of the Pur box. Binding of the helix-turn-helix motif is then allowed at the major grove, and, altogether, transcription is impaired (104, 256, 534, 535). Results of analysis of purR mutant proteins have confirmed a role of Lys54 in DNA bending, and a role of Lys55 in binding to the adenine 5' to the central CpG (22, 177). The purine repressor is also involved in the expression of operons other than pur and purR. These include pyrC, pyrD, and codBA of pyrimidine metabolism, the prs gene, the glyA gene, the gcv operon, as well as speA and glnB. The effect of purR on these various pathways has been reviewed previously (681).
IMP formed by the de novo biosynthetic pathway is converted to AMP via succinyl-AMP and to GMP via XMP. AMP and GMP are then phosphorylated to ADP and GDP by specific nucleoside monophosphate kinases and further converted to the triphosphate level as described in a previous section. The biosynthesis of AMP from IMP occurs with the intermediate formation of succinyl-AMP and requires the participation of adenylosuccinate synthase, encoded by purA and adenylosuccinate lyase, encoded by purB (Fig. 5, Table 3). The use of GTP as the energy donor for the adenylosuccinate synthase reaction, feedback inhibition by AMP, a 2- to 3-fold repression by PurR, and an uncharacterized adenine-dependent repression mechanism (218) balance the production of adenine nucleotides to that of guanine nucleotides.
The biosynthesis of GMP from IMP requires the participation of two enzymes, IMP dehydrogenase (encoded by guaB), which forms XMP and GMP synthase (encoded by guaA), which replaces the exocyclic oxygen at the 2 position of XMP with an amino group (Fig. 5). The genes are organized in the guaBA operon (331) and transcribed from a single promoter, which is repressed approximately 5-fold by PurR in response to the presence of guanine or hypoxanthine (397). Activity of the promoter is also subject to a 15-fold regulation by the DNA initiator protein DnaA, which binds to two DnaA boxes about 200 bp apart (576). An AT-rich UP-element in the region −59 to −38 interacting with C-terminal domain of the α-subunit of RNA polymerase is necessary for the high level of transcription and involved in the growth-rate-dependent control of guaBA transcription (126, 258). The use of ATP as the energy donor in these reactions probably contributes to balancing the production of guanine nucleotides to the adenine nucleotide pools.
Individual enzyme reactions.
In solution, the enzyme is a mixture of 47-kDa monomers and dimers with a dissociation constant for the dimer, KD = 10 μM. The dimer is greatly stabilized by the substrate IMP (and analogues), because residues from both subunits contribute to substrate binding in a shared active site (31, 302, 633). The reaction kinetics and the catalytic mechanism of the E. coli enzyme have been studied in detail and a large number of crystal structures of wild-type and mutant enzymes in complexes with substrate analogues and inhibitors have been determined, previously reviewed, and discussed in connection with the reaction mechanism (238, 239). In short, the Mg2+-requiring reaction, which follows rapid equilibrium kinetics, involves two chemical steps: (i) the transfer of the γ-phosphate of GTP to IMP, forming a 6-phosphoryl-IMP intermediate and (ii) the subsequent nucleophilic displacement of phosphate from P-IMP by the α-amino group of aspartate to generate succinyl-AMP. The existence of the phosphorylated intermediate, first predicted on the basis of the enzyme-catalyzed, IMP-dependent rearrangement of β,γ-bridging 18O-oxygen of GTP (30), was later confirmed by crystallographic studies (102, 483). The enzyme is inhibited by IMP and AMP, which may be regarded as a feedback inhibitor, by the aspartate analogues succinate and Hadacidin, and by GDP and ppGpp, which bind in the GTP site and may have a physiological function during nutritional shifts (94, 241). A mathematical model accounting for the reaction kinetics of this enzyme has been described (466).
The use of glutamine is eliminated by alkylation of Cys86, which is situated in the glutaminase domain, but the alkylated enzyme is still active with ammonia. On the other hand, all enzyme activity is eliminated by reaction with the glutamine analogue 6-diazo-5-oxonorleucine (679, 683) as is characteristic of enzymes using glutamine as amide donor (682). GMP synthesis proceeds via an enzyme-bound intermediate O2-adenyl-XMP, which by reaction with ammonia generates the products GMP and AMP (622). The enzyme is a tetramer of 525 residue subunits (217, 522, 523). The tetramer is arranged as a dimer of dimers. The glutaminase activity resides in a class 1 amidotransferase domain, which is formed by the N-terminal 206 amino acid residues of each subunit and contains a catalytic triad, Cys86, His181, and Glu183, similar to the catalytic triads of cysteine proteinases. The residues 207 to 406 constitute the synthetase domain, and the residual C-terminal part of the polypeptide is responsible for dimer formation (577). There are no reports of allosteric control of enzyme activity.
The pathways for salvage and interconversion of adenine and guanine compounds are shown in Fig. 5. In total, these pathways ensure that mutants unable to synthesize IMP by the de novo pathway can satisfy the purine requirement by any purine base or nucleoside present in the growth medium.
Individual enzyme reactions.
The enzyme is essential for purine-requiring mutants grown with guanine or xanthine as sole sources of purine (Fig. 5). The KM for GMP is about 0.1 mM (378). ATP inhibits the enzyme and the inhibition is fully reverted by GTP (458). Transcription of the gene may be under nitrogen control (315) and is induced by high concentrations of guanosine in the culture medium (42). The enzyme is a tetramer composed of four identical subunits with a molecular weight of 37 kDa (17). No crystal structure of a GMP reductase from enteric bacteria is available, but the enzyme may resemble the human enzyme (PDB ID code 2BLE; unpublished).
The histidine biosynthetic pathway produces one molecule of AICAR (and hence IMP) per turn and contributes, therefore, to the interconversion of adenine and 6-oxopurine nucleotides. The enzyme belongs to the type I PRTases with a classical PRTase fold and forms hexamers in the crystals (368) and solution (6, 620). The reaction is inhibited by AMP, which binds in the PRPP site in such a way that the ATP site becomes blocked by the adenine ring of AMP (368). Histidine serves as a feedback inhibitor and rearranges the oligomeric state of the enzyme (6, 368, 620) perhaps in a synergistic fashion with ppGpp, which accumulates during nutritional downshifts (414). Enzyme synthesis is regulated by attenuation in a leader peptide region, which contains seven histidine codons in front of a Rho-independent transcription terminator and is thus repressed at a high charging level of histidinyl-tRNA (29, 89).
De novo synthesis of the pyrimidine nucleoside triphosphates, UTP and CTP, proceeds by a linear pathway initiated by the formation of carbamoyl phosphate from the phosphate (and energy) of ATP, bicarbonate, and ammonia from glutamine. The aromatic pyrimidine base orotate is formed in three steps and subsequently equipped with a phosphoribosyl group resulting in the formation of the first pyrimidine nucleotide orotidine 5'-monophosphate (OMP). UMP is made by decarboxylation of OMP and converted to UTP by two subsequent kinase reactions. Finally, CTP is made from UTP by exchange of the 4-OH group with an amino group. The pathway is illustrated in Fig. 6. In the following section we will discuss each of these reactions individually.
Enzymes of pyrimidine de novo synthesis.
The enzyme can also use ammonia from solution in place of that derived from glutamine hydrolysis (370).
The enzyme is regulated allosterically by GTP, which stimulates the glutaminase-dependent reaction at concentrations up to 0.15 mM in vitro. Above this concentration, inhibition of both the glutamine-dependent and the ammonia-dependent CTP synthesis reaction occurs (375, 376, 653). However, the glutaminase reaction appears to be uncoupled from CTP formation under these conditions because the hydrolysis of glutamine is unaffected at GTP concentrations up to at least 0.5 mM (376). Apart from stimulating the glutaminase reaction, GTP seems somehow to close the tunnel that channels nascent ammonia between the two active sites (268, 374). Although the GTP binding site has yet not been identified in the crystal structure of the enzyme, several structure–function studies have supported a binding site between the two-subunit domains as suggested by modeling studies (141). The triphosphate chain interacts with residues in the glutamine amidotransferase domain to activate hydrolysis of glutamine (32, 375, 654, 659), whereas the nucleoside moiety of GTP is suggested to close the ammonia channel (375). The activation by GTP seems to be of physiological importance also at concentrations higher than 0.15 mM GTP, because, in growing cells, reduction of the GTP pool from about 1 mM to about 0.2 mM causes reduction of the CTP pool and a small increase in the UTP pool (269, 273, 491).
The product CTP inhibits the enzyme by binding in the active site at a position slightly different but overlapping that proposed for UTP and the newly synthesized CTP (142). In this way CTP also acts as an allosteric inhibitor. CTP synthase is unique by binding UTP, CTP, and ATP as substrates or products and CTP and GTP as effectors whereby all ribonucleotides have influence on the activity of the enzyme.
Individual genes and operons.
The de novo pathway for pyrimidine nucleotide synthesis generates both UTP and CTP. The CTP synthase reaction is the only way to form cytidine nucleotides and cytidine/cytosine compounds can only be converted to uracil compounds by the deamination of cytosine or cytidine (437) (see Fig. 6). The nucleoside monophosphate kinases, UMP kinase and CMP kinase, in combination with nucleoside diphosphate kinase reconvert the pyrimidine nucleotide products from RNA breakdown to diphosphates and triphosphates. While UMP kinase is an integral part of the de novo pathway leading to UTP (via UDP) and CTP, the CMP kinase only plays a role in salvage of RNA breakdown products. This salvage reaction is of major importance for formation of CDP, which seems to be the major pyrimidine substrate for ribonucleotide reduction under aerobic growth conditions, since bacteria lacking CMP kinase have reduced pools of dTTP and dCTP and exhibit reduced rates of DNA chain elongation (158). Exogenous pyrimidine nucleosides and bases are good sources for intracellular nucleotide synthesis and are taken up by specific transport systems as described in a subsequent section. Cytosine and cytidine are generally deaminated to the corresponding uracil compounds before they are converted to UMP either by uracil PRTase or uridine/cytidine kinase. Deamination to the corresponding uracil compounds is the only way to utilize cytosine and deoxycytidine for nucleotide synthesis, because no organism is known contain a cytosine PRTase or a (reversible) cytidine phosphorylase and enteric bacteria do not have a deoxycytidine kinase (308, 437). Bacteria can also use exogenous uridine and cytidine nucleotides for endogenous nucleotide synthesis, but this ability depends on the specific growth conditions, because the nucleotides must be dephosphorylated before they are taken up as nucleosides, and this ability depends on the level of alkaline phosphatase, which is repressed by high phosphate concentrations in the medium (60, 437). Orotate is taken up by the dicarboxylic acid transport system (27) and is converted to OMP by orotate PRTase. It is a good exogenous pyrimidine source for auxotrophic mutants in minimal medium, but supports only slow growth in the presence of the amino acid mixtures that contain aspartate and glutamate (618).
Individual enzyme reactions.
Deoxyribonucleotides are made de novo by reduction of ribonucleoside di- or triphosphates catalyzed by a ribonucleotide reductase. When formed the deoxyribonucleoside diphosphates are phosphorylated to the triphosphates by nucleoside diphosphate kinase. The three products, dATP, dGTP, and dCTP, are used for DNA synthesis. The fourth substrate dTTP used for DNA synthesis is formed in a series of reactions by which dUTP is first converted to dUMP by dUTPase and then methylated at the 5 position by thymidylate synthase to form dTMP, which finally is converted to dTDP by thymidylate kinase and further to dTTP by nucleoside diphosphate kinase. Most of the dUTP (about 75%) used for dTTP synthesis is made by deamination of dCTP, catalyzed by dCTP deaminase rather than by being generated directly from UDP by the action of ribonucleotide reductase and nucleoside diphosphate kinase. The pathways are shown in Fig. 7, which also illustrates the tight regulation of the activity of the ribonucleotide reductases by feedback and feed-forward mechanisms that ensure a balanced supply of deoxyribonucleotides under different growth conditions. dTMP (and thus dTTP) can also be made through salvage reactions by phosphorylation of thymidine, catalyzed by thymidine kinase and from thymine by reversal of the thymidine phosphorylase reaction when the intracellular pool of the cosubstrate deoxyribose 1-phosphate is sufficiently high as it is in thymine-requiring (thyA) mutants. The source of deoxyribose 1-phosphate is the very high pool of dUMP that accumulates in such strains. This nucleotide is dephosphorylated to deoxyuridine and then phosphorolytically cleaved to yield uracil and deoxyribose 1-phosphate. The different enzymatic steps in these pathways are described in detail below; the properties of the genes and enzymes are compiled in Table 5.
The ribonucleotide reductase operating under exponential and aerobic growth conditions in both E. coli and S. enterica is NrdAB (a class Ia enzyme), which is encoded by the nrdAB genes (209). The class 1b ribonucleotide reductase, NrdEF, is encoded by the nrdHIEF operon, which in addition to the two ribonucleotide reductase subunits also codes for an electron donor protein, NrdH, and a protein of unknown function (292). A single copy of the nrdHIEF operon is unable to support growth of an nrdAB strain, probably because NrdR very strongly represses nrdHIEF operon expression, but nrdAB strains are viable if additional copies of the nrdHIEF operon are present either on the chromosome or on plasmids (180, 293). In both E. coli and S. enterica the expression of the nrdAB operon is controlled in a complex manner by regulatory proteins involved in cell-cycle control and growth-phase regulation, i.e., DnaA, Fis (23, 180, 294), and IciA (206). In addition, expression of both the nrdAB and nrdHIEF operons is increased in response to DNA damage or inhibition of DNA synthesis (172, 407). Expression of the nrdHIEF operon is strongly increased by exposure of the cell to oxidative stress and in mutants lacking enzymes involved in the breakdown of peroxides and superoxides (407). However, the regulatory mechanism of elevated nrdHIEF expression under these conditions is still unknown and appears to be different from the known major regulatory mechanisms involved in oxidative stress (407). A FUR box upstream of the nrdHIEF operon seems to play a role in the regulation of expression in response to iron limitation as demonstrated by global transcription analysis of a fur mutant (615).
The crystal structures of representative ribonucleotide reductases from class Ia (E. coli [604]), class Ib (S. enterica [605, 606]), and anaerobic class III (T4 phage [336]) are available (Table 5). Both class Ia, structurally very similar class Ib, and class III ribonucleotide reductases are heterotetramers composed of two heterodimers that are organized either with a symmetry axis perpendicular to (class Ia and Ib) or in the direction of the dimer interface (class III). The heterodimer consists of an R1 (or α) and an R2 (or β) subunit where the active site, the effector sites, and specificity sites are on the R1 subunit. The nrdA, nrdE and nrdB, nrdF genes of the two operons specifying the class I ribonucleotide reductases encode the two subunits R1 and R2, respectively. The folding topology of the R1 subunit, an α/β barrel, of all three classes of ribonucleotide reductases is conserved together with critical active-site residues. The binding site for the specificity effectors, which dictate what ribonucleotide substrate should be reduced, is situated at the R1 subunit interface. The initial formation of the important radical that is shuffled to the active site of R1 is formed in the R2 subunit of the class I ribonucleotide reductases that contains the diiron site and the tyrosyl radical. The radical is transferred through the enzyme to a cysteine in the large R1 subunit that activates the substrate. The catalytic mechanism involves first the formation of thiyl radical that activates the substrate at C-3' by abstracting a proton and forming a substrate radical that subsequently leads to formation of a double bond between the C-3' and C-2' and elimination of the 2'-hydroxyl. For the class Ia enzyme the next round of the catalytic cycle proceeds by donation of an electron from thioredoxin or glutaredoxin to reduce the keto-tautomer of the substrate at C-3'. For the nrdEF encoded class Ib ribonucleotide reductase the redox protein is the NrdH protein encoded by the same operon as the catalytic subunits (137, 321, 452).
In the class III ribonucleotide reductase the R1 and R2 subunits are encoded by the nrdD and nrdG genes. For this enzyme that is active under anaerobic conditions the reduced iron-sulfur cluster of the R2 subunit is believed to cleave S-adenosylmethionine to form a 5'-deoxyadenosyl radical that through a glycyl radical intermediate forms the thiyl radical at the active site of the R1 subunit (15).
The class I enzymes reduce ribonucleoside diphosphates, whereas the class III enzyme reduces ribonucleotide triphosphates. The specificity regulation of substrate reduction for both class I (139, 333, 334) and class III (140) ribonucleotide reductases are as follows: dATP and, in class Ia enzyme, also ATP, stimulates reduction of CDP (CTP) and UDP (UTP), dTTP stimulates reduction of GDP (GTP), and dGTP stimulates reduction of ADP (ATP). In addition, class Ia and class III enzymes have a regulatory site, the activity site, where binding of ATP or dATP, respectively, activates or inhibits the overall activity of the enzyme (140, 333, 334). The importance of the allosteric regulation of ribonucleotide reductase seems unquestionable for its role in maintaining stable deoxyribonucleotide pools and the impact of these on genome stability (386, 650).
dCTP deaminase and dUTPase.
The dUDP produced by reduction of UDP becomes phosphorylated by nucleoside diphosphate kinase and cleaved by dUTPase to form diphosphate and dUMP, which is used for dTMP synthesis. An alternative pathway for supplying dUMP is the deamination of dCTP catalyzed by dCTP deaminase followed by hydrolysis of dUTP to dUMP and diphosphate catalyzed by dUTPase. The dCTP deaminase pathway contributes about 75% of the total dUMP for dTMP synthesis both in E. coli (460) and S. enterica (436, 444). The importance of the dCTP pathway is illustrated by the observation that a dcd mutant, devoid of dCTP deaminase activity, has a 10-fold increased dCTP pool and a 4-fold reduced dTTP pool, which become (partly) normalized by addition of thymidine to the medium (443). A dcd insertion mutation was also found in a selection for mutants having a constitutive SOS phenotype following random transposon mutagenesis (461), indicating that, even though cells lacking dCTP deaminase activity grow normally and have no easily detected phenotype (443), they are stressed by the imbalance of the dCTP and dTTP pools. Recently, a third pathway for formation of dUMP has been revealed, which most likely plays a much less significant role in wild-type cells. This pathway involves the dephosphorylation of dCMP, arising primarily from phospholipid metabolism, by the 5'-nucleotidase encoded by the ybfR gene followed by deamination of deoxycytidine and phosphorylation of deoxyuridine to dUMP by thymidine kinase (640, 641).
Whereas dUTPase binds dUTP very specifically, dCTP deaminase binds both dCTP and dUTP. In addition, dCTP deaminase binds the inhibitor dTTP, which is the ultimate end product of the pathway initiated by this enzyme (34, 290). The dTTP regulation is mediated by its binding to an inactive form of dCTP deaminase. Binding of a single dTTP molecule to one active site alters the position of the catalytic side chains in all three active sites due to a steric clash between the methyl group of the thymine moiety and an alanine residue that, in turn, forces a local rearrangement of the peptide backbone. It has been shown that the transition between the active and inactive forms of dCTP deaminase is a slow process that, at least in part, can account for the increased cooperativity of dCTP saturation induced by dTTP (290).
dTMP synthase or thymidylate synthase.
The structure of thymidylate synthase has been solved in complex with 5-fluoro-dUMP and the folate substrate, N5,N10-methylene-THF, and analogues thereof (259, 388, 389). The enzyme is a dimer where the interface between the subunits is formed by the central six-stranded β-sheet of each subunit. The subunit shows an active-site cleft that extends some 25 Å of the interior of the protein. The formation of the ternary complex results in major structural changes where the C-terminal closes down over the folate substrate and bound nucleotide to shield the active site from the solvent.
dTMP kinase.
Thymidylate kinase was described in the section on nucleoside monophosphate kinases, above.
Thymidine kinase.
Thymidine phosphorylase.
but is inactive with ribonucleosides. The reaction is readily reversible, but the equilibrium favors nucleoside formation. The enzyme is responsible for the ability of thyA mutant strains to use thymine as source of dTTP and it requires a high intracellular concentration of deoxyribose 1-phosphate. As described above, this happens in thyA strains, but wild-type strains do not incorporate exogenous thymine in DNA due to shortage of deoxyribose 1-phosphate unless other deoxyribonucleosides are present in the medium (280). Thymidine phosphorylase from both E. coli and S. enterica has been purified and characterized in detail (54, 232, 536, 537). The reaction mechanism is sequential with phosphate as the first substrate to bind and deoxyribose 1-phosphate the last product to leave (537). Unlike the other nucleoside phosphorylases, thymidine phosphorylase is a homodimer; its structure is very different from those of uridine phosphorylase and the purine nucleoside phosphorylases. Each subunit is composed of a small α-helical domain of six helices and a large α/β domain, and the active site lies in a cavity between the small and large domains. Movements of the domains relative to each other may explain the kinetic reaction order (494, 627).
5'-Nucleotidase.
Two 5'-nucleotidases, YfbR and YjjG, which are potentially relevant for the dephosphorylation of dUMP, were identified by screening crude extracts for general phosphatase activity and subsequently characterizing the substrate specificity of the purified enzymes (493). YfbR is a Co2+-dependent phosphatase, which shows a broad specificity for 5'-deoxyribonucleoside monophosphates, but is inactive with the corresponding ribonucleotides. YjjG requires Mg2+ and has a narrow specificity for 5'-dTMP, 5'-dUMP, and 5'-UMP (493). The YjjG nucleotidase has been identified as the enzyme responsible for the physiological dephosphorylation of dUMP, since thyA yjjG mutants are unable to use thymine as external source of dTTP (641). Crystal structures of the YfbR nucleotidase have been determined (689).
Tsx.
NupC.
NupG.
XapB.
NepI.
YegT.
YegT is a low-affinity purine-specific transporter that is 28% identical to NupG and XapB and belongs to the MFS family (B. Mygind and G. Dandanell, unpublished data). YegT is predicted to consist of 12 TM segments with the C terminus inside (118).
UraA.
CodB.
PurP.
YgfO and YicE.
YgfO and YicE are homologs (45% identity) that belong to the NCS2 family. Both are high-affinity, xanthine-specific transporters (KM 3 to 4 μM) (188, 304). YgfO is predicted to consist of 12 TM segments with the C terminus inside (118). Substrate binding studies using purine analogues and cysteine scanning experiments indicate that xanthine binds the transporter via hydrogen bonds with the basic interactions to the pyrimidine ring (N3-H and O-2) and the imidazole ring (N-9) (188, 305).
Other transporters.
Regulation of the genes encoding nucleobase transporters has not been studied in detail, but none of the characterized transporters appears to be regulated by DeoR or CytR.
Xanthosine uptake and salvage is not coregulated with the other salvage enzymes and transporters. However, induction of xapAB depends on the presence of active NupC and NupG. Unlike the other genes in nucleoside salvage, which are always expressed at a low basal level, the xapAB operon is only expressed when E. coli is grown in the presence of xanthosine or deoxyinosine (201, 297). Transcription of xapAB is activated in the presence of inducer by its specific LysR-type activator XapR, whose gene is located downstream of xapAB (540). Although XapB is the primary xanthosine transporter, the basal level of XapB is too low to provide sufficient amount of xanthosine required to induce transcription of xapAB. NupC or NupG are required to take up sufficient amounts of xanthosine to induce transcription of xapAB (201, 540). S. enterica cannot utilize xanthosine as carbon source. All necessary genes appear to be present, but a Gly72Asp alteration in XapA of S. enterica serovar Typhimurium (but not in other serovars) prevents the utilization of xanthosine (211). Induction of XapA expression has been proposed to be under catabolite repression, but no CRP binding site has been found and such catabolite repression probably does not involve CRP (84). Xanthosine can also be hydrolyzed by RihC. However, since a xapA mutant (expressing xapB) cannot grow on xanthosine, the expression of rihC may be too low to enable E. coli and S. enterica to utilize xanthosine as carbon source (210, 211, 478). Expression of rihC is under catabolite repression (478).
Cytidine is catabolized by deamination to uridine, which can be phosphorolyzed to uracil and ribose 1-phosphate by cytidine deaminase and uridine phosphorylase, respectively. Transcription of both cdd and udp is induced by cytidine by relieving the repression by CytR. As other CytR-regulated genes are, they are both under catabolite repression by CRP-cAMP (70, 200, 205, 236, 298, 611). The udp gene as well as cdd is expressed from a single promoter where a single CytR molecule binds between two CRP molecules. CRP-cAMP acts both as activator and corepressor for CytR (611, 690).
The pathways for degradation of pyrimidine ribo- and deoxyribonucleotides to the free pyrimidine bases and the pentose moieties have been known for many years. Ribose 1-phosphate and deoxyribose 1-phosphate, which arise from phosphorolytic cleavage of the nucleosides, are converted to derivatives of central metabolism by participation of enzymes of the deo-operon (204), but pathways for degradation of the pyrimidine ring in E. coli have remained obscure. Many bacteria degrade the pyrimidine bases by a reductive pathway, which includes (i) the reduction of the pyrimidine bases uracil and thymine to the corresponding dihydropyrimidine compounds, catalyzed by the enzyme dihydropyrimidine dehydrogenase (EC 1.3.1.2), followed by (ii) an opening of the ring structure, catalyzed by the enzyme dihydropyrimidinase (EC 3.5.2.2), and finally by (iii) breakdown of the product N-carbamoyl-β-alanine (or, in the case of thymine, N-carbamoyl-β-isobutyric acid), catalyzed by the enzyme N-carbamoyl-β-alanine amidohydrolase (EC 3.5.1.6), yielding β-alanine (619). This pathway seems to be operative in E. coli B (648) although the identity of the genes specifying the enzymes, especially that catalyzing the last step, iii, is somewhat hypothetical. E. coli K-12 and S. enterica are unable to use uracil as the sole nitrogen source at 37°C, although the organisms do possess genes for the pathway enzymes. However, recently it was found that E. coli was able to degrade uracil and thymine by a hitherto unrecognized catabolic pathway, which is active at 25°C, but not at 37°C, and which is present only during nitrogen limitation (367). The pathway does not conform to the reductive or oxidative pathways, known from other organisms, but leads to the release of two molecules of ammonia and 3-hydroxypropionic acid (lactic acid) or 2-methyl-3-hydroxypropionic acid from uracil or thymine, respectively. The individual reaction steps of the pathway are still unknown, but seven enzymes encoded by the rutABCDEFG operon are all required for catabolism of these pyrimidines. Transcription of the operon is under nitrogen control and requires the participation of NtrC. A gene rutR, encoding a repressor RutR of the operon is transcribed from the promoter region in the opposite direction—away from the catabolic operon genes—by the σ70 RNA polymerase (367). The catabolic operon genes rutABCDEFG are absent from the chromosomes of all Salmonella species, but interestingly the rutR gene is retained (545). The RutR protein belongs to the TetR repressor family and the structure of it was determined some years ago prior to knowledge of its function (PDB ID code 1pb6). A search for binding sites for RutR on the E. coli chromosome was carried out by a SELEX technique and several sites were identified: a site in the rut-operon promoter region, a site in the rutR promoter, indicating autoregulation of rutR expression, sites in promoter regions of genes controlling glutamate and glutamine metabolism and allantoin degradation, and most interestingly a site in the carAB P1-promoter region, overlapping one of the two binding sites for the PepA repressor (545). Foot-printing and band-shift assays further identified uracil (and to a lesser extent thymine) as the effector of the RutR protein, causing its release from the DNA binding sites (545). In this way the presence of uracil both leads to induction of the synthesis of enzymes needed for its degradation when the nitrogen supply is poor, and repression of pathways for de novo synthesis of pyrimidine nucleotides and arginine by liberating the site for PepA repressor binding to the carAB P1 promoter (545).
Based on sequence similarity it was suggested that the open reading frames b2866, b2867, and b2868 in E. coli might encode molybdate binding, FAD binding, and FeS binding subunits, respectively, of a putative xanthine dehydrogenase, and since production of a minute amount of 14CO2 from uniformly labeled 14C-adenine, seen in the wild-type strain was eliminated by deletion of the b2866, the reading frames were renamed xdhA, xdhB, and xdhC and it was proposed that E. coli could oxidize the purine ring to uric acid by means of xanthine dehydrogenase and further to allantoin by a putative uricase (668). However, the employed E. coli strain could not utilize the purine ring-atoms as nitrogen source and a potential gene for the putative uricase was not identified (668). The xdhABC genes have no homologs in Salmonella species or in Klebsiella species that are known to degrade the purine ring. However, E. coli is capable of using allantoin, but not the intact purine ring as nitrogen during anaerobic growth, and degrading it to oxaluric acid (117). Genes encoding proteins with similarity to the needed enzymes: allantoinase (EC 3.5.2.5, P77671, 49.6 kDa, allB, b0512), allantoate amidohydrolase (EC 3.5.3.4, P77425, 45.7 kDa, allC, b0516), ureidoglycolate hydrolase (EC 3.5.3.19, P77731, 18.2 kDa, allA, b0505), and ureidoglycolate dehydrogenase (EC 1.1.1.114, P77555, 40.0 kDa, allD, b0517) were cloned, expressed, and shown to have the predicted enzymatic activities (117). These genes are also present in S. enterica: allB (STM0523), allC (STM0527), allA (STM0515), and allD (STM0528) and in K. pneumoniae.
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Module3.6.2
