Biosynthesis and Conversions of Pyrimidines
Chapter
35
JAN NEUHARD and ROD A. KELLN
Intracellular pyrimidine compounds are almost exclusively found as nucleotides; i.e., the N-1 position of the nucleobase is linked by a β-N-glycosyl to either d-ribose phosphate or its 2-deoxyribose derivative. More than 95% of the nucleotides are found in the acid-insoluble fraction of the cells as nucleic acids. The acid-soluble fraction consists of nucleoside mono-, di-, and triphosphates (NMPs, NDPs, and NTPs) and nucleotide-containing coenzymes. NTPs are the immediate precursors of both the nucleic acids and the coenzymes. The nucleotide-containing coenzymes, as well as a small fraction of the nucleic acids, i.e., the mRNAs, undergo a high rate of turnover, yielding NMPs and NDPs as products. For stable RNA and DNA synthesis, a constant supply of ribonucleoside triphosphates (rNTPs) and deoxyribonucleoside triphosphates (dNTPs) is required, which in the absence of preformed utilizable precursors must be provided through de novo biosynthesis.
The pathway for de novo synthesis of the pyrimidine NTPs may be regarded as an unbranched sequence of enzyme reactions in which dTTP is the ultimate end product and UTP, CTP, and dCTP are obligatory intermediates (Fig. 1). The product of the first reaction, carbamoylphosphate, is also an intermediate of arginine biosynthesis and constitutes the only branch point of the pathway. To provide the cells with a balanced supply of rNTPs and dNTPs for RNA and DNA synthesis, the pathway is regulated through allosteric enzymes at five strategic points (Fig. 1): (i) the first reaction of the pathway, which is the synthesis of carbamoylphosphate; (ii) the first reaction specific to pyrimidine nucleotide synthesis, which is the formation of carbamoylaspartate; (iii) the amination of UTP to form CTP; (iv) the reduction of CTP to dCTP; and (v) the conversion of dCTP to dTTP. In addition, the pathway is controlled at the level of gene expression by mechanisms regulating the synthesis of the six enzymes catalyzing the de novo synthesis of UMP and of ribonucleotide reductase, the enzyme specific for the key crossover reaction from ribonucleotides to deoxyribonucleotides. Certain intermediates of the pathway can be derived directly from preformed pyrimidine compounds present in the growth medium via the salvage pathways, through which exogenous pyrimidine bases and nucleosides are converted to intracellular pyrimidine NMPs.
The pathway responsible for UTP and CTP biosynthesis is shown in Fig. 2. It is initiated by the formation of carbamoylphosphate from glutamine, bicarbonate, and ATP, a reaction catalyzed by carbamoylphosphate synthetase (CPSase) encoded by the carAB operon. Aspartate carbamoyltransferase (ATCase), a multimeric enzyme encoded by the pyrBI operon, catalyzes the first unique reaction of pyrimidine biosynthesis, namely, the condensation of carbamoylphosphate and the amino group of aspartate to form carbamoylaspartate (ureidosuccinate). The reaction is driven by the high-energy acid anhydride bond of carbamoylphosphate. In the next two steps, the pyrimidine ring is formed through cyclization and oxidation. The first reaction is catalyzed by dihydroorotase (DHOase) (pyrC) and results in ring closure to yield dihydroorotate. Dihydroorotate dehydrogenase (DHOdehase) (pyrD), which is membrane bound and linked to the electron transport system, carries out the oxidation of dihydroorotate to orotate. The first pyrimidine nucleotide is formed by the transfer of ribose 5-phosphate from α-d-5-phosphoribosyl-1-pyrophosphate (PRPP) to orotate, forming orotidine 5'-monophosphate (OMP), which is subsequently decarboxylated to UMP. The reactions are catalyzed by orotate phosphoribosyltransferase (OPRTase) (pyrE) and OMP decarboxylase (OMPdecase) (pyrF), respectively.
The generation of UTP involves phosphorylation of UMP to UDP and then to UTP by the sequential action of UMP kinase (pyrH) and NDP kinase (NDK) (ndk). Finally, UTP is aminated to CTP by CTP synthetase (pyrG). The enzyme uses glutamine as the preferred amino group donor and ATP to energize the reaction. Although CMP is not an obligatory intermediate in de novo CTP synthesis, Escherichia coli and Salmonella typhimurium (official designation, Salmonella enterica serovar Typhimurium) each have a CMP kinase (cmk) distinct from UMP kinase; the enzyme also uses dCMP as substrate. Presumably, the physiological function of this enzyme is to rephosphorylate the CMP produced by the turnover of mRNA and CDP-diacylglycerols (23, 70a).
CPSase (carAB).
CPSase (EC 6.3.5.5) is an allosteric enzyme that catalyzes the synthesis of carbamoylphosphate from bicarbonate and glutamine at the expense of two molecules of ATP. One ATP is required for the formation of the reaction intermediate, carboxyphosphate, and the other is required for a kinase reaction in which enzyme-bound carbamate becomes phosphorylated (11). The enzyme also catalyzes carbamoylphosphate synthesis with ammonia replacing glutamine, but the affinity for ammonia is much lower than that for glutamine.
The kinetic properties of CPSases from E. coli (145, 219, 220) and S. typhimurium (4) have been studied. The Mg-ATP saturation curves are sigmoidal, whereas Michaelis-Menten kinetics are observed for glutamine and bicarbonate. Activity is inhibited by UMP and activated by ornithine, IMP, and, for the Salmonella enzyme, PRPP. The effectors influence activity by altering the affinity of the enzyme for Mg-ATP. The enzyme contains one binding site for each of the effectors (ornithine, IMP, and UMP) (9); the binding sites for UMP and IMP overlap (32). Thus, CPSase is feedback regulated in accordance with its metabolic role. If the supply of carbamoylphosphate becomes limiting for arginine synthesis, ornithine accumulates and antagonizes the inhibition by UMP. In the presence of excess arginine, ornithine is not produced, and the enzyme is controlled solely by UMP. The physiological importance of this control is illustrated by the observation that mutations rendering CPSase hypersensitive to inhibition by UMP induce a uracil sensitivity phenotype (174).
The CPSase holoenzyme (160 kDa) is a heterodimer consisting of a glutamine-binding subunit with glutaminase activity (42 kDa; carA) and a synthetase subunit (118 kDa; carB) (Table 1). The holoenzyme can be reversibly dissociated into its requisite small and large subunits. The synthetase subunit carries the binding sites for the substrates bicarbonate and ATP and can effect carbamoylphosphate synthesis from bicarbonate, ATP, and ammonia. It exhibits a sigmoidal saturation curve for Mg-ATP and also contains the binding sites for the effectors (145, 192). Two separate and functionally different ATP binding sites have been identified; one is in the amino-terminal half, and the other is in the carboxy-terminal half of the subunit (33, 178). The binding site for UMP is also located in the carboxy-terminal domain (192). The primary amino acid sequence of the large subunit exhibits a highly significant homology between the amino- and carboxy-terminal halves, suggesting that the carB gene may have evolved from a smaller ancestral gene by duplication (167). On the basis of amino acid sequence similarities between the small subunit and other glutamine amidotransferases and of studies of mutant enzymes altered in the putative glutaminase domain, the carboxy-terminal half of the small subunit is judged to be responsible for the hydrolysis and channeling of the amide-N of glutamine to the catalytic center of the synthetase subunit (117, 176, 191). The N-terminal third of the small subunit is essential for interaction with the large subunit (78).
Table 1Properties of pyrimidine biosynthetic enzymes and corresponding genes |
ATCase (pyrBI).
Since the pioneering work of Gerhart and Pardee (see reference 71), ATCase (EC 2.1.3.2; also called aspartate transcarbamoylase) from E. coli has become one of the most extensively studied allosteric enzymes (91, 107, 108, 109, 198, 199). It catalyzes the first unique step of pyrimidine nucleotide biosynthesis whereby carbamoylphosphate is reacted with l-aspartate to form N-carbamoyl-l-aspartate and Pi. The enzyme is activated by ATP and feedback inhibited by CTP; ATP decreases the apparent Km of the enzyme for aspartate, and CTP increases it. UTP alone does not inhibit activity, but in the presence of CTP, a synergistic inhibition occurs (64, 65, 247, 256).
The ATCase holoenzyme (310 kDa) is a dodecamer composed of two catalytic trimers (c3; 102 kDa) and three regulatory dimers (r2; 34 kDa) (Table 1). The holoenzyme shows a sigmoidal saturation curve for both substrates and therefore exhibits positive homotropic cooperativity. Heterotropic effects on the activity of the enzyme are induced by the nucleotide effectors. The cooperative homotropic and heterotropic interactions reflect a conformational transition of the holoenzyme between a less-active low-affinity state, the T state, and a more-active high-affinity state, the R state, and involve intricate protein-protein interactions between both homologous chains (c-c and r-r interactions) and heterologous chains (c-r interactions) (41, 53, 107, 172, 210, 211).
Treatment with mercurials or heat dissociates the holoenzyme into catalytic (c3) and regulatory (r2) subunits. The catalytic subunit is enzymatically active but lacks the homotropic response to the substrates (i.e., it exhibits hyperbolic kinetics), and it is insensitive to inhibition by CTP or activation by ATP. The regulatory subunit is catalytically inactive but binds the nucleotide effectors competitively to the same site. The regulatory subunit contains a zinc-binding domain (140), and in the presence of zinc ions, a fully active holoenzyme showing normal allosteric behavior can be formed from the isolated subunits (36, 143, 155).
The three-dimensional arrangement of the catalytic and regulatory subunits of the holoenzyme was determined by X-ray crystallography, and this approach was applied to probe the changes in quaternary structure occurring upon binding of the substrates or substrate analogs, or nucleotide effectors (75, 76, 113, 122, 210). Chemically or mutationally modified ATCases were used to study the structural basis of catalysis and allosteric regulation (53, 61, 96, 120, 163, 172, 242, 251, 257). A molecular modeling study on the mechanism of catalysis was reported (74), and a number of mechanisms were suggested to account for heterotropic regulation (211).
ATCase from S. typhimurium appears to be very similar to the E. coli enzyme both in structure and in catalytic and regulatory properties (169, 246). Intergeneric hybrid enzymes composed of E. coli catalytic subunits with S. typhimurium regulatory subunits, and vice versa, show cooperative saturation curves for aspartate.
DHOase (pyrC).
DHOase (EC 3.5.2.3) is a homodimer with a subunit molecular weight of 38 kDa (Table 1). It catalyzes the cyclization of N-carbamoyl-l-aspartate, the product of the ATCase reaction, to l-dihydroorotate. The reaction is readily reversible, the amide bond formation being favored at pHs below 7. The E. coli enzyme was purified to homogeneity; it contains one tightly bound essential zinc ion per subunit (240). In addition, it has two weakly bound structural zinc ions per subunit, but they are not essential for activity (241). The enzyme is sensitive to oxygen in the presence of trace metal ions. Comparison of the amino acid sequences of DHOases from various species reveals very little sequence identity (182), but two conserved histidine-containing regions that are likely to be involved in active-site zinc binding and in catalysis are present (40).
DHOdehase (pyrD).
The only redox reaction in de novo UMP biosynthesis is the oxidation of dihydroorotate to orotate, a reaction catalyzed by the membrane-bound DHOdehase (EC 1.3.3.1; also called dihydroorotate oxidase; 110). There is evidence that the physiological electron acceptor in the presence of oxygen is ubiquinone (116); in anaerobically grown cells, menaquinone appears to be an obligatory hydrogen carrier between dihydroorotate and fumarate (12). The homodimeric enzyme (subunit size, 37 kDa; Table 1) was purified to near homogeneity from membranes of E. coli (111, 123). The purified enzyme cannot use oxygen but can use ferricyanide or 2,6-dichlorophenol-indophenol as an electron acceptor. The absorption spectrum of the oxidized enzyme indicates one flavin nucleotide per subunit. The addition of dihydroorotate in the absence of an electron acceptor results in bleaching of the spectrum, indicating that the bound flavin is a cofactor in the reaction; the flavin nucleotide has been identified as flavin mononucleotide (123). The amino acid sequences of a number of DHOdehases contain a highly conserved region that resembles the cofactor binding site of flavoproteins (123, 153). This conserved region is present in the S. typhimurium enzyme as well (70).
OPRTase (pyrE).
The nucleotide-forming step in the pathway consists of the Mg2+-dependent formation of OMP and PPi from orotate and PRPP, a reaction catalyzed by OPRTase (EC 2.2.4.10; also called OMP pyrophosphorylase). The sole function of Mg2+ in the reaction is formation of a monomagnesium complex with the PPi group of PRPP, Mg2+-PRPP, which is the actual ribosyl donor in the reaction (27). The reaction is accompanied by anomeric inversion at C-1 of the ribosyl group to give a β-N-glycosyl. The active form of OPRTase is a homodimer formed from 23-kDa subunits (Table 1). The enzyme from both organisms has been crystallized (5, 197), and the structure of the S. typhimurium enzyme complexed with OMP has been determined (196). Kinetic studies with pure S. typhimurium OPRTase indicated a random sequential mechanism with no ping-pong properties (28). Amino acid sequence alignments of a number of OPRTases revealed a conserved sequence also found in a number of other phosphoribosyltransferases and in PRPP synthetase and inferred to be the PRPP binding site (95, 182). The large number of negative charges associated with the substrates and products suggests the involvement of basic amino acid residues in the active site. On the basis of chemical modification of S. typhimurium OPRTase, three specific lysines have been indicated as active-site residues (77); all three are conserved in those OPRTases for which sequence data are available.
OMPdecase (pyrF).
The final reaction of de novo pyrimidine nucleotide biosynthesis is catalyzed by OMPdecase (EC 4.1.1.23), whereby OMP is irreversibly decarboxylated to yield UMP, the parent compound for all other pyrimidine nucleotides. The decarboxylation proceeds in the absence of a coenzyme requirement, unlike amino acid decarboxylation or the decarboxylation of pyruvate, and the reaction may occur via a noncovalent zwitterionic intermediate that is readily decarboxyated to form an ylide (dipolar carbanion). After decarboxylation, proton transfer yields the uracil moiety of the UMP (20). The analog, 6-azauridine 5'-monophosphate, is a competitive inhibitor (84). The subunit molecular mass is 26 kDa (Table 1; 217, 225), and the enzyme appears to be catalytically active as a dimer (59, 104). Alignment of the amino acid sequences with those of 18 other enzymes (both prokaryotic and eukaryotic) revealed four conserved sequences that were present in all 20 polypeptides (119).
UMP Kinase (pyrH) and CMP Kinase (cmk).
Studies with mutants defective in UMP kinase (EC 2.7.4.?) and CMP kinase (EC 2.7.4.14) activity established that these two activities are catalyzed by distinct enzymes (23, 97). The nucleotide sequence of the E. coli pyrH gene (204) is identical to that of smbA (252), a gene originally identified through its involvement in suppression of a null mutation in mukB, which encodes a protein required for proper chromosomal partitioning. The pyrH gene product has a deduced subunit molecular mass of 26 kDa (Table 1). Purified UMP kinase is active as a homohexamer and is specific for UMP; dUMP, CMP, and dCMP are not substrates. UMP kinase is allosterically regulated by nucleotides, with GTP a positive effector and UTP a negative effector (202a). CMP kinase lacks specificity toward the sugar and uses both CMP and dCMP as substrates. cmk null mutants display a decreased replication elongation rate, presumably as a result of the reduction of the intracellular dCTP and dTTP pools, but this decreased rate is compensated for by a doubling in the frequency of initiations (70a). Thus, rephosphorylation of CMP arising from the turnover of mRNA and CDP-diacylglycerols may contribute significantly to the synthesis of CDP and thereby of dCTP and dTTP (see Deoxyribonucleotide Biosynthesis below). The cmk gene has been cloned as a multicopy suppressor gene, mssA, of the conditional lethal phenotype of certain smbA (pyrH) mutants (251a).
NDK (ndk).
The last reaction in the synthesis of NTPs is catalyzed by NDK (EC 2.7.4.6), an enzyme with very little specificity toward the base or sugar moiety. All rNTPs and dNTPs can function as donors of the γ-phosphate group, and any rNDP or dNDP may serve as the phosphate acceptor (73, 170). Mg2+ is required for activity, and the reaction occurs in two steps, with the intermediate formation of a phosphoenzyme(73, 80). The deduced molecular mass of the E. coli K-12 NDK subunit is 15.5 kDa, which is in agreement with the subunit molecular mass for the purified enzyme (16 kDa; Table 1). The active form of the enzyme appears to be a tetramer, i.e., 64 kDa (170). However, the reported molecular masses of partially purified NDKs from various strains of E. coli vary: for E. coli B, it is 55 kDa (195), and for E. coli K108, it is 115 kDa (186). The E. coli K108 enzyme is loosely bound to the outer surface of the cytoplasmic membrane, a location that seems incompatible with the generation of intracellular NTPs. The amino acid sequence of E. coli NDK exhibits 42 to 57% identity with the sequences of six other NDKs (80).
CTP Synthetase (pyrG).
The last step in pyrimidine ribonucleotide biosynthesis consists of the formation of CTP from UTP, ATP, and glutamine, a reaction catalyzed by CTP synthetase (EC 6.3.4.2) and stimulated by GTP (121). Like other glutamine amidotransferases, CTP synthetase can utilize ammonia instead of glutamine, although with much lower efficiency. The molecular mass of the E. coli CTP synthetase subunit is 60 kDa (Table 1; 244). The enzyme was purified to homogeneity, and its physical and kinetic properties were described (10, 121). Under assay conditions, the enzyme exists as a homotetramer, but in the absence of ATP and UTP, it dissociates to dimers. The reaction mechanism involves the initial glutamylation of a specific cysteine residue on the enzyme, with the liberation of ammonia. The nascent ammonia immediately reacts with UTP, and in a series of steps that probably involves a phosphorylated amido derivative of UTP (229a), CTP is formed at the expense of one ATP. GTP markedly accelerates the glutamine reaction by increasing the V max and decreasing the Km for glutamine, but it has no effect on the ammonia reaction. Amino acid sequence alignments with other glutamine amidotransferases identified the active-site cysteine within a glutamine amide transfer domain located in the carboxy-terminal half of the polypeptide (244). The enzyme exhibits negative cooperativity for the effector GTP and the substrate glutamine and pronounced positive cooperativity for the substrates ATP and UTP. The conformational changes induced by ATP and UTP result in conversion of the enzyme from a dimer to a tetramer. In addition, ATP and UTP appear to be allosteric effectors of the glutamylation reaction, indicating the induction of conformational changes affecting the glutamine amide transfer domain.
S. typhimurium mutants and, with the exception of ndk mutants, E. coli mutants defective in each of the nine steps to CTP synthesis have been isolated and characterized. The genetic loci specifying the enzymes have been mapped and shown to be well separated on the chromosomes. The locations of the E. coli genes on the physical map of the chromosome are known, and the E. coli genes and most of the S. typhimurium genes have been cloned and had their sequences determined (Table 1). Four of the six enzymes arise from operons: the carAB operon encodes the two subunits of CPSase (117, 167, 176), and the polypeptides of native ATCase are encoded by the pyrBI operon (147, 200); the pyrE gene is the distal gene of a dicistronic operon, rph-pyrE (162, 179, 181, 197), in which rph specifies RNase PH (171); pyrF is the promoter proximal gene of the pyrF-orfF operon (217, 225), but the functional identity of the orfF gene product has yet to be determined. The remaining two enzymes are encoded by the individual pyrC (16, 160, 249) and pyrD (70, 123) genes.
Null mutations in carA and carB produce a dual requirement for arginine and a pyrimidine in accordance with the function of CPSase in supplying carbamoylphosphate for both arginine and pyrimidine biosynthesis. However, a number of different phenotypes are found among CPSase mutants (3, 146). Certain missense mutations cause auxotrophy only for arginine, a phenotype that is frequently more pronounced at low temperatures (2, 83, 146). Studies of such cold-sensitive S. typhimurium mutants indicated that the conditional arginine requirement resulted from defective folding of CPSase at the nonpermissive temperature (83). Uracil or arginine sensitivity may also result from mutations in carAB. One uracil-sensitive mutation causes hypersensitivity of CPSase to feedback inhibition by UMP (174). The biochemical basis for the arginine sensitivity phenotype has been studied in S. typhimurium (1). Ornithine carbamoyltransferase, an enzyme of arginine biosynthesis, appears to be required for proper assembly of the subunits of the mutant enzyme. Thus, arginine inhibits growth by repressing the synthesis of ornithine carbamoyltransferase. Whether ornithine carbamoyltransferase is also involved in the assembly of the wild-type carbamoylphosphate synthetase is not known.
Mutations in pyrBI, pyrC, pyrD, pyrE, or pyrF typically result in pyrimidine auxotrophy. Wild-type cells are not permeable to the intermediates of the pathway except for orotate. Orotate satisfies the pyrimidine requirement of carAB, pyrB, pyrC, and pyrD mutants, provided glycerol is used as carbon source (253). With glucose as carbon source, orotate is growth rate limiting for carAB and pyrB mutants in a concentration-dependent manner (234). Carbamoylaspartate (ureidosuccinate)-permeable mutants (usp) have been isolated (126, 216), and a pyrB usp double mutant grows on carbamoylaspartate as the sole pyrimidine source. Partial pyrimidine starvation of S. typhimurium pyrC and pyrD mutants results in severe growth inhibition due to intracellular accumulation of carbamoylaspartate, but the mechanism of this inhibition has not been identified (223). Certain strains of E. coli K-12, e.g., W3110 and derivatives, excrete orotate into the growth medium (135). This excretion can be explained by the observation that each of these strains contains a frameshift mutation in the rph gene of the rph-pyrE operon, leading to reduced synthesis of the OPRTase (102).
Expression of the genes and small operons encoding the first six enzymes of the pathway is regulated in a complex manner by the intracellular nucleotide pools, involving both pyrimidine and purine nucleotides and, for the carAB operon, arginine. The nucleotide effectors responsible for the regulation were identified largely through the use of mutant strains that allowed manipulation of individual nucleotide pools in vivo (Table 2; 101, 114, 133, 173, 202). Expression of the genes is regulated noncoordinately, with some genes responding to changes in the UTP pool and others responding to fluctuations in the CTP pool. However, mutants exhibiting a simultaneous increase in the expression of several pyr genes have been characterized. In most cases, this phenotype was caused by mutations in genes encoding enzymes involved in nucleotide interconversion, resulting in altered effector nucleotide pools. These include bradytrophic pyrH mutants defective in UMP kinase (97, 106, 173) and bradytrophic guaB mutants with low IMP dehydrogenase activity (99). In mutants in which altered pyr gene regulation did not result from disturbed nucleotide pools, the mutations were localized in rpoBC, implying a direct involvement of RNA polymerase in the regulation of pyr gene expression (105, 158). Superimposed on the nucleotide control, a twofold repression of carAB, pyrC, and pyrD expression is mediated by purine bases. This control requires a functional purine repressor, which is encoded by purR. In the presence of one of the corepressors, guanine or hypoxanthine, PurR binds to a purine operator sequence located in the promoter regions of these genes (49, 161, 230, 250).
Table 2Expression of genes of de novo pyrimidine biosynthesis |
carAB
.
The synthesis of CPSase is subject to cumulative repression by arginine and pyrimidines (54, 117, 176). A repressive effect of purines on carAB expression has also been observed (133; C.-D. Lu and A. T. Abdelal, personal communication). A direct role for RNA polymerase in the regulatory mechanism stems from the isolation of S. typhimurium rpo mutants exhibiting altered pyrimidine regulation of CPSase synthesis (105, 158).
The carAB operon is transcribed from tandem promoters (P1 and P2) 67 bp apart that are controlled by pyrimidines and arginine, respectively. The escape from both arginine and pyrimidine control under multicopy conditions suggests the involvement of titratable regulatory elements in both control systems (117). The P2 promoter overlaps one of two adjacent 18-bp sequences, ARG boxes, which are characteristic of operators of arg genes. Repression of P2 expression requires arginine-dependent binding of the hexameric arginine repressor (the argR gene product) to both of the adjacent ARG boxes (46, 47, 132, 190). The combination of pyrimidines and arginine represses P2 more efficiently than arginine alone and is dependent on a functional P1 promoter; this effect is not the result of pyrimidine binding to ArgR (47, 132).
Expression from P1 is regulated negatively by pyrimidines, with both uracil and cytosine nucleotides being effective (54, 117, 133, 173). A number of trans-acting factors appear to be involved in the control of transcriptional initiation from P1. In both E. coli and S. typhimurium, integration host factor (IHF) binds specifically to a 38-bp AT-rich region positioned 305 bp upstream of the P1 transcriptional start point, resulting in enhanced expression from P1 in minimal medium and reduced expression in pyrimidine-supplemented medium (45). In E. coli, dominant mutations in carP, a locus unlinked to carAB, result in constitutive expression from P1 (190). The nucleotide sequence of the carP gene is identical to that of xerB, which encodes a component of the system for resolving ColE1 multimers into monomers, and to pepA, which encodes aminopeptidase A. The purified carP-xerB-pepA gene product, a hexamer of 55-kDa subunits, binds in vitro to two 25-bp sites located 67 bp apart between the IHF binding site and the P1 promoter. This binding is independent of the presence of pyrimidines (D. Charlier, personal communication). Protection in vivo of a GATC Dam methylation site 106 bp upstream of the P1 transcriptional start site is dependent on high pyrimidine nucleotide pools and on binding of both IHF and CarP to their respective target sites (44, 238).
In S. typhimurium, the use-1 mutation imparts temperature-sensitive hyperrepression of carAB expression by pyrimidines (uracil sensitivity; 42), and this effect is exerted at P1 (117). The use gene has been identified as an allele of argU, which encodes a tRNA for a minor arginine codon; the use-1 mutation is a base substitution in the anticodon stem of the tRNA (131). The basis for the uracil sensitivity phenotype is not understood, but involvement of a translational event in pyrimidine control of carAB expression is likely.
Purines negatively control carAB expression from the P1 promoter. The purine holorepressor binds to a purine operator sequence (PUR box) located at position –128 relative to the P1 transcriptional start point (Lu and Abdelal, personal communication). This binding further emphasizes the importance of the nontranscribed region upstream of P1 as a target region for regulatory factors controlling transcription. In addition, transcription from P1 appears to be subject to stringent control, indicating an added involvement of purines (ppGpp) in expression (37, 176).
pyrBI
and pyrE.
Expression of pyrBI and pyrE is regulated at the transcriptional level, and the synthesis of ATCase and OPRTase is repressed when the intracellular UTP pool is increased and derepressed when it is lowered (101, 114, 202). An inverse correlation between synthesis and the size of the GTP pool has also been observed (99, 101, 180).
For both organisms, the DNA upstream of the start of the pyrB (147, 154, 189, 224) and pyrE (162, 179) genes is very similarly structured and contains a rho-independent transcriptional terminator sequence (attenuator). Another region of dyad symmetry (transcriptional pause site), followed by a sequence encoding a uridylate-rich cluster in the transcript, precedes the attenuator. The leader regions of the mRNAs are open reading frames; for pyrBI, the leader encodes a short polypeptide of 44 (E. coli) or 33 (S. typhimurium) amino acids, whereas for pyrE, the leader is the rph mRNA that codes for the RNase PH polypeptide. The essential features of attenuation control require that transcription terminate at the attenuator in the absence of closely coupled translation and that termination be prevented if transcription and translation are coupled. Tight coupling between the transcribing RNA polymerase and the first ribosome translating the leader mRNA is required for transcription through the attenuator. When the UTP concentration is high, transcription and translation are effectively uncoupled, facilitating termination at the attenuator. When the UTP concentration is low, however, this polarity is relieved by the reduced rate of elongation by RNA polymerase through the uridylate-rich pause region, which allows for coupling with the more rapidly progressing lead ribosome.
Direct evidence for involvement of the attenuator in regulation was obtained by showing that chromosomal mutations leading to elevated expression can result from base substitutions in the attenuator region that reduce the stability of the putative hairpin structure of the transcript (159). Furthermore, plasmid constructs deleted for the attenuator region have elevated expression and reduced repressibilty (17, 128, 130).
The necessity of translation of the leader region for expression and UTP-mediated regulation has been demonstrated for both pyrBI (50, 187, 188) and pyrE (35, 179). Moreover, the partial pyrimidine requirement observed for certain E. coli K-12 "wild-type" strains, namely, W3110 and MG1655, results from a frameshift mutation in rph that arises through the deletion of a G-C base pair near the 3' end and leads to the termination of polypeptide synthesis 10 codons in advance of the normal termination site (102). In S. typhimurium, insertion of Mud1 in rph results in an absolute pyrimidine requirement (162). Translation through the attenuator is, however, not a prerequisite for preventing transcriptional termination (35, 147, 187); for pyrBI, the ribosome must translate to within 14 to 16 nucleotides of the attenuator-encoded leader mRNA hairpin to inhibit transcriptional termination efficiently (187).
Alteration of the rate of transcription or translation can affect expression of the two operons. In vitro transcription analyses revealed the presence of NusA-dependent pause sites in rph, including a major pause site after the first region of dyad symmetry, and expression of pyrE in a nusA mutant was hyperrepressed (6). Correlatively, a decrease in the rate of transcription imparted by a mutant RNA polymerase with decreased affinity for UTP resulted in constitutively derepressed expression of pyrBI and pyrE (103, 105). In contrast, a decrease in the rate of translation impaired derepression (100), but increasing the rate of ribosomal movement across the terminal region of rph resulted in an increased level of expression under repressing conditions (34).
The frequency of initiation of transcription for rph-pyrE is virtually independent of the intracellular UTP concentration (179), whereas a 6- to 10-fold effect on E. coli pyrBI expression has been reported (57, 127, 130); such additional regulation does not appear to occur for S. typhimurium pyrBI (17). Since control through attenuation is influenced by the rate of mRNA chain elongation, the fact that fluctuations in the GTP pool also impact on the expression of the two operons has been interpreted on the basis of kinetic properties of elongating RNA polymerase (101). A further involvement of guanine nucleotides in expression stems from the observations that accumulation of ppGpp inhibits ATCase synthesis in vivo (222) and that ppGpp can inhibit transcription from the E. coli pyrBI promoter in vitro (57).
pyrC
and pyrD.
Expression of pyrC and pyrD increases during CTP starvation but decreases when the GTP pool decreases (101, 114, 173, 202). For both organisms, the leader regions of pyrC (16, 160, 249) and pyrD (70, 123) are short (36 and 33 bp, repectively) and nontranslated and do not contain sequences indicating the presence of an attenuator. However, the leaders contain a region of hyphenated dyad symmetry, which may allow the 5' end of the transcript to form a stable secondary structure that sequesters sequences necessary for ribosomal binding and which was the basis for forwarding a regulatory model involving translational attenuation (115). The importance of the symmetry region in the regulation of pyrC and pyrD was demonstrated by the finding that point mutations, which destabilize the putative secondary structure in the leader transcript, result in high-level constitutive expression (70, 115, 248). Determination of the 5' ends of the pyrC and pyrD transcripts revealed nucleotide pool-dependent heterogeneity in the selection of the transcriptional initiation site within the start site motifs (CCGG for pyrC and CCCGG for pyrD) located 7 and 6 bp, respectively, downstream of the –10 region (70, 207, 208, 248).
At high CTP/GTP intracellular pool ratios, i.e., repressing conditions, transcription starts with the first C within the start site motif, producing an mRNA that is inefficiently translated because of its capacity to form the stable hairpin at the 5' end (Fig. 3). At a low CTP/GTP pool ratio, the predominant start is not the first C but occurs with the first G, 2 or 3 bp further downstream. This shorter transcript has a reduced potential to form the stable secondary structure at the 5' end, and translation is unhindered. Direct evidence for the presence or absence of the proposed secondary structure in the pyrC and pyrD transcripts was obtained by chemical and enzymatic probing of total RNA from repressed and derepressed S. typhimurium cultures (206).
Addition of purines to the growth medium leads to a twofold repression of DHOase and DHOdehase synthesis. The PurR holorepressor binds to the PUR box sequence located in the promoter regulatory region; in pyrC, the PUR box is located between the –35 and –10 regions, whereas in pyrD, it is found 90 bp upstream of the transcriptional start point (49, 161, 230, 250). Thus, two independent purine-mediated control systems, apparently operating in opposition to one another, are involved in regulating pyrC and pyrD expression. However, regulation through translational attenuation resulting from transcriptional start site variation is not structured to impose purine control but rather to facilitate pyrimidine control by sensing fluctuations in the relative intracellular CTP concentration. Direct purine control is limited to the repressive action of hypoxanthine and guanine through interaction with the purine repressor.
pyrF
.
Expression of the pyrF-orfF operon is negatively regulated 6- to 10-fold by a uracil nucleotide (114, 202) and 3-fold by a guanine nucleotide (99); expression of pyrF and orfF is translationally coupled (218). The pyrF leader region does not exhibit the characteristic features inherent in the leaders of the pyrBI and rph-pyrE operons, and it therefore appears that a control mechanism based on UTP-modulated attenuation is not operative for this operon. Primer extension analyses (225) and studies with pyrF-galK operon fusions (218) indicate that expression is regulated at the transcriptional level. Results obtained from measuring the rate of pyrF mRNA synthesis in vivo in conjunction with determinations of the level of OMPdecase activity led to a proposal that mRNA half-life rather than mRNA synthesis per se may be the mode by which pyrimidine-control is exerted (M. Theisen, Ph.D. thesis, University of Copenhagen, Copenhagen, Denmark, 1989). The finding that a strain deficient in polynucleotide phosphorylase has a higher level of OMPdecase than its isogenic control (58) supports this notion.
The pyrimidine salvage pathways of E. coli and S. typhimurium (Fig. 4) serve three physiological functions. The first is assimilation of exogenous free bases and nucleosides; the nucleosides are predominantly metabolized to free bases before being used for nucleotide synthesis. The second function is making the pentose moieties of exogenous nucleosides available as a source of carbon and energy (see chapter 20 of this volume) and the amino groups of cytosine compounds available as a nitrogen source. The third function is reutilizing free bases and nucleosides produced intracellularly from nucleotide turnover. Significant amounts of ribonucleotides are degraded during normal growth, and the reutilization of these free bases and nucleosides requires salvage enzymes (82, 166, 175).
Cytotoxic pyrimidine analogs, including the 5-fluoropyrimidine bases and nucleosides, are toxic after their conversion to nucleotides. Such analogs have been used extensively to select for mutants defective in salvage pathway enzymes. Studies of these mutants have contributed significantly to establishing the physiological functions of the individual enzymes. Table 3 summarizes the phenotypes of certain mutants in which salvage is affected (see also reference 157).
Table 3Effects of mutations in pyrimidine salvage pathways on utilization of pyrimidines and sensitivity toward 5-fluoropyrimidines |
Uracil.
Uracil is converted to UMP primarily by uracil phosphoribosyltransferase (UPRTase) (upp). Alternatively, in the presence of high intracellular concentrations of one of the substrates for uridine phosphorylase (udp), that is, uracil or ribose 1-phosphate, uracil may be converted to UMP through the concerted action of uridine phosphorylase and uridine kinase.
UPRTase (EC 2.4.2.9), which is specified by upp, catalyzes the Mg2+-dependent conversion of uracil and PRPP to UMP and PPi. The homotrimeric enzyme from E. coli (subunit size, 23 kDa) is activated by GTP (Table 4; 66, 183). The upp gene is the first gene of a bicistronic operon in which the second gene, uraA, encodes a cytoplasmic membrane protein required for uracil uptake (7a). Pyrimidine starvation results in increased levels of UPRTase, indicating that expression is under pyrimidine control (8, 183). Mutants defective in UPRTase excrete uracil, showing that free uracil is produced endogenously, presumably from nucleotide turnover (175).
Table 4Properties of enzymes and genes of pyrimidine salvage pathways |
Cytosine.
The only known route for cytosine metabolism is through deamination to uracil and ammonia. Cytosine deaminase (EC 3.5.4.1), which is encoded by codA, was purified to homogeneity from both E. coli (112, 177) and S. typhimurium (245) and appears to be a homotetramer of 48-kDa subunits. The E. coli enzyme contains one catalytically essential divalent metal ion per subunit and is most active with Fe2+ (Table 4; 177). The codA gene is the second gene of a bicistronic operon in which the first gene, codB, encodes a permease for cytosine (see discussion on transport below; 56). Expression of the operon is controlled in a complex manner; expression is repressed by purines and derepressed by starvation for either pyrimidines or nitrogen (7). The purine holorepressor binds to a purine operator sequence located between the –35 and –10 regions of the cod promoter. Nitrogen control is mediated by the glnLG (ntrBC) gene products (7, 118). Thus, the cod operon belongs to both the nitrogen regulatory system (NTR) and the PurR regulons.
Uridine.
Uridine is converted to UMP either by phosphorylation catalyzed by uridine kinase (udk) or by phosphorolytic cleavage to uracil and ribose 1-phosphate through the action of uridine phosphorylase (udp) and then conversion of the uracil to UMP via UPRTase (see above). Uridine kinase (EC 2.7.1.48) is specific for ribonucleosides (deoxyribonucleosides are not substrates) and catalyzes the phosphorylation of uridine and cytidine to UMP and CMP, respectively, with GTP (or dGTP) as the preferred phosphate donor; UTP and CTP inhibit the reaction (Table 4; 227). Gel filtration analysis of partially purified E. coli uridine kinase indicated an approximate molecular mass of 90 kDa. With a deduced subunit size of 23 kDa, the enzyme appears to be a homotetramer. Expression of udk is derepressed during pyrimidine starvation, but the repressing nucleotide has not been identified.
Uridine phosphorylase (EC 2.4.2.3) catalyzes the reversible phosphorolysis of uridine. It shows weak activity with deoxyuridine and thymidine as substrates but no activity toward cytidine or deoxycytidine (Table 4) (125, 232). The E. coli enzyme was purified to homogeneity and crystallized (52, 125, 233). Chemical modification studies provided evidence for the involvement of a histidine residue in the active site of the enzyme (60). There have been conflicting reports on the quaternary structure of the enzyme, but the crystal structure indicates that the enzyme is a hexamer of 27-kDa subunits (52, 236). The synthesis of uridine phosphorylase is coregulated with the synthesis of a number of proteins involved in nucleoside transport and catabolism. Thus, expression of the udp and cdd genes, the deo operon (see chapter 20 of this volume), and the genetic loci for three nucleoside transport systems (see below) is negatively controlled by the cytR-specified repressor and positively controlled by the cyclic AMP (cAMP) receptor protein (CRP)-cAMP complex. The inducer of the CytR regulon in E. coli is cytidine, whereas both cytidine and uridine are effectors in S. typhimurium (81, 166, 205). Measurements of the growth yield from uridine of pyrimidine-requiring mutants defective in the ability to utilize uracil (upp mutants) established that a large fraction (50 to 75%) of the added uridine becomes unavailable for the kinase reaction as a result of phosphorolysis to uracil (157).
Cytidine.
The predominant route for cytidine utilization is through deamination to uridine catalyzed by the inducible cytidine deaminase (cdd; Table 4). Any cytidine that escapes deamination may be converted directly to CMP by uridine kinase (see above). Cytidine deaminase (EC 3.5.4.5) catalyzes the deamination of both cytidine and deoxycytidine as well as their 5-methyl and 5-fluoro derivatives to the corresponding uracil nucleosides. The V max/Km ratio for deoxycytidine is 10-fold higher than that for cytidine (13, 94, 231). The enzyme from E. coli consists of two identical subunits (31.5 kDa each), each containing one tightly bound zinc atom (254). Characterization of transition-state analog inhibitors of the enzyme supports a proposed reaction mechanism involving direct water attack at the C-4 atom of the cytosine ring (14, 51, 69). Further evidence for this mechanism is derived from X-ray crystallographic studies of a cytidine deaminase–transition-state analog complex in which a zinc atom in the active site is complexed to two cysteines and one histidine ligand (26).
Mutants defective in CTP synthetase, i.e., pyrG mutants, require cytidine for growth. However, this growth requirement cannot be met unless the mutants contain a cdd mutation rendering them defective in cytidine deamination (22). Pyrimidine-auxotrophic cdd mutants are able to grow, albeit slowly, on cytidine as the sole pyrimidine source. This residual growth is mediated by an enzyme activity that catalyzes the hydrolysis of CMP to cytosine and ribose 5-phosphate (A. Eisenhardt, M.Sc. thesis, University of Copenhagen, Copenhagen, Denmark, 1971). Accordingly, such growth is abolished by mutations inactivating either cytosine deaminase (codA) or uridine (cytidine) kinase (udk; Table 3). As a component of the CytR regulon, expression of cdd is induced by cytidine (uridine) and involves both the cytR-encoded repressor and the CRP-cAMP complex as described above for udp expression (228).
Deoxyuridine and Thymidine.
Deoxyuridine and thymidine can also be utilized by two different routes. Thymidine kinase, which is encoded by tdk (31), phosphorylates either compound to the monophosphate level. In a second route, thymidine phosphorylase (deoA) cleaves these nucleosides to deoxyribose 1-phosphate and the corresponding free base. Thymidine kinase (EC 2.7.1.21; Table 4) from E. coli is an allosteric enzyme that is feedback inhibited by dTTP and activated by a number of dNDPs and dNTPs, with dCTP being the most effective (48). The native enzyme appears to be active as a homotetramer of 23.5-kDa monomers (31). Mutants deficent in thymidine kinase cannot use thymidine or deoxyuridine as specific precursors for DNA synthesis.
Thymidine phosphorylase (EC 2.4.2.4) catalyzes the reversible phosphorolysis of pyrimidine deoxyribonucleosides except for 4-amino-substituted compounds such as deoxycytidine. Ribonucleosides are not cleaved, and uridine or ribose 1-phosphate inhibits activity (157). The enzyme from both organisms is a homodimer with a subunit molecular mass of 45 kDa (Table 4; 235). The three-dimensional structure of the E. coli enzyme has been determined, and the active site has been identified (235). Mutants defective in thymidine phosphorylase can still utilize deoxyuridine as the sole pyrimidine source, whereas deoA udp double mutants cannot. Thus, uridine phosphorylase also degrades deoxyuridine to some extent in vivo. The deoA gene is part of the deo operon, and consequently, synthesis of thymidine phosphorylase is negatively controlled by the repressor proteins specified by deoR and cytR and positively controlled by the CRP-cAMP complex; both deoxyribonucleosides and cytidine (uridine) induce expression (55, 81, 205; see chapter 20 of this volume).
Deoxycytidine.
E. coli and S. typhimurium are devoid of deoxycytidine kinase activity and are also incapable of directly converting deoxycytidine to cytosine and a deoxyribose compound. However, deoxycytidine can be effectively deaminated to deoxyuridine, and this conversion requires a functional cytidine deaminase. Thus, deoxycytidine is not metabolized by cdd mutants.
Pyrimidine ribo- and deoxyribonucleotides serve as the sole pyrimidine sources in both E. coli and S. typhimurium (129, 226). Their utilization is dependent on a number of periplasmic phosphatases that hydrolyze nucleotides to nucleosides (157, 226). The nucleosides are subsequently taken up via specific transport systems located in the cytoplasmic membrane (see below). In addition, specific outer membrane pore-forming proteins are required for passage of the nucleotides into the periplasmic space (19, 229).
Bases.
Primarily on the basis of genetic evidence, specific transport systems for uracil and cytosine have been identified in E. coli (152). Mutants defective in uracil uptake contain mutations in uraA, the distal gene of the upp-uraA operon. The uraA gene product (45 kDa) is located in the cytoplasmic membrane, and the deduced amino acid sequence indicates that the protein may be organized in the membrane and have 12 transmembrane α-helices (7a). A pyrimidine-requiring uraA mutant grows normally with cytosine as the sole pyrimidine source, whereas the growth rate on uracil at concentrations below 80 μM is strictly dependent on the uracil concentration. At low uracil concentrations, uracil uptake requires both the uraA and the upp gene products, suggesting that such uptake occurs by facilitated diffusion mediated by UraA followed by conversion to UMP by UPRTase.
Cytosine transport requires the cytoplasmic membrane protein cytosine permease (subunit M r, 42 kDa), which is encoded by codB, the first gene of the codBA operon (56). The transport is active, and the system is capable of concentrating extracellular cytosine about 200-fold. The topology of the codB gene product in the membrane has been studied by using codB::phoA fusions, and the protein appears to be organized with 12 transmembrane α-helices (S. Danielsen, personal communication). The cod operon is regulated at the transcriptional level by purines, nitrogen, and pyrimidines, as discussed above.
Nucleosides.
Two high-affinity active-transport systems for nucleosides, the C and G systems, have been characterized in E. coli (152). The G system transports all common ribo- and deoxyribonucleosides and is inactivated by mutations in nupG. The C system, specified by nupC, is limited to the transport of pyrimidine and adenine nucleosides. Both nupC and nupG encode a 45-kDa polypeptide, but the two polypeptides exhibit no significant amino acid sequence similarity (53a, 87); NupG and NupC are localized within the cytoplasmic membrane. A double mutant (nupC nupG) cannot utilize nucleosides as a source of carbon and energy, but the ability of pyrimidine nucleosides to satisify a pyrimidine requirement is not fully abolished.
The E. coli outer membrane protein Tsx, the product of the tsx gene, functions as a nucleoside-specific channel as well as a receptor for colicin K and a number of T-even bacteriophages such as T6 (88, 144, 201). The Tsx polypeptide (31 kDa) shows no significant sequence similarity to other channel-forming proteins (39). Mutation of tsx impairs the transport of all nucleosides except cytidine and deoxycytidine.
Synthesis of the two transport systems and Tsx is coregulated with the production of enzymes for nucleoside catabolism. Expression of nupC, nupG, and tsx is regulated by the cytR-encoded repressor and the CRP-cAMP complex; nupG and tsx are also controlled by the deoR-encoded repressor (39, 72, 151).
Deoxyribonucleotides are synthesized from their corresponding ribonucleotides by enzymes termed ribonucleotide reductases (Fig. 5). In the reaction, the C-2 hydroxyl of the d-ribose is replaced by a hydrogen, yielding deoxyribose. Three classes of ribonucleotide reductases have been identified, and certain properties are shared among them: (i) they reduce all four common ribonucleotides, (ii) their activities and substrate specificities are allosterically controlled by ATP and dNTPs, and (iii) they may utilize a free-radical amino acid residue in the reduction process (185, 213). Class I is typified by the E. coli rNDP reductase (RDPR) and contains an oxygen-linked iron (Fe3+) center. Class II enzymes reduce rNTPs and require deoxyadenosyl cobalamin for the reaction; no enzyme belonging to this class has been detected in E. coli or S. typhimurium. Class III, found in anaerobically grown E. coli, also reduces rNTPs, but requires S-adenosylmethionine (AdoMet) as coenzyme.
RDPR (nrdAB).
RDPR (EC 1.17.4.1), a class I enzyme from aerobically grown E. coli, has been studied extensively (68, 184, 185, 212, 213). It has the general tetrameric protein structure, α 2 β 2 (Table 5), and consists of component proteins R1 (the α 2 dimer of 85.7-kDa protomers; formerly called B1) and R2 (β 2; 43-kDa protomers; formerly called B2). The crystal structures of the R1 protein (226a) and R2 protein (165) have been determined. Each R1 subunit contains a substrate binding site with a redox-active sulfhydryl pair and two independent effector binding sites that control both catalytic activity and substrate specificity. The diferric iron center (Fe3+-O-Fe3+) of each of the R2 subunits functions in generation and maintenance of an essential tyrosyl radical located on Tyr-122 (124, 165). The enzyme is inhibited by radical scavengers such as hydroxyurea and by 8-hydroxyquinoline, which chelates the iron, resulting in simultaneous loss of the radical.
Table 5Properties of enzymes and genes of deoxynucleotide metabolism |
The first step of the proposed catalytic sequence (185, 212, 213) involves abstraction of the hydrogen atom attached to C-3'. Initially, the tyrosyl radical was thought to have a direct involvement, but the current view is that a thiyl radical of Cys-439 may be directly involved, with the tyrosyl radical having the role of a radical initiator (138, 165). Abstraction of the hydrogen atom is followed by loss of a hydroxide ion from C-2', which then becomes reduced through thiols of the enzyme (137). The hydrogen atom abstracted from C-3' is transferred back to its original position to complete the transformation of the rNDP to a dNDP.
In allosteric regulation, both the activity and the specificity of the enzyme are regulated by NTP effectors through binding to specific allosteric sites on the R1 protein. The activity site binds ATP and dATP; with ATP bound, the enzyme is active, but when dATP is bound, the enzyme is inactive. The substrate specificity site binds ATP, dATP, dTTP, and dGTP, and the nature of the bound effector determines which rNDP will be the preferred substrate (185).
Evidence for a new bacterial ribonucleotide reductase in both S. typhimurium and E. coli has been obtained (105a). In both organisms, the nrdEF operon encoding the new reductase is located just upstream of proU, near 57 min on the chromosome. The deduced amino acid sequences of the nrdE and nrdF products include the catalytically important residues conserved in the class I ribonucleotide reductases.
rNTP Reductase (nrdD).
The class I enzyme could be expected to be rendered nonfunctional in anaerobic conditions, since oxygen is needed for generation of the tyrosyl radical (184). Anaerobically grown E. coli contains an oxygen-sensitive rNTP reductase, a class III enzyme, that requires AdoMet for activity (90). The reductase system consists of three protein components, with one component (dA3), which is encoded by nrdD (215), being the reductase per se (63). One of the other components (dA1) is ferredoxin (flavodoxin) NADP+ reductase (frp; 29), and the last component is presumably either flavodoxin or ferredoxin. The enzyme is an iron-sulfur protein structured as a 160-kDa homodimer (150, 215). Upon initial isolation, it is inactive; activation requires anaerobic preincubation with NADPH, AdoMet, and the other two protein components. Activation may involve the production of an oxygen-sensitive radical of a glycine residue near the carboxy terminus, and NADPH, AdoMet, and the Fe-S cluster may take part in this process (185). The protein shows 72% amino acid sequence identity with the polypeptide encoded by the bacteriophage T4 sunY gene, a gene with a self-splicing intron and no currently known function (255).
In allosteric regulation, substrate specificity is governed by specific effectors, and 10-fold stimulation of activity can be achieved with appropriate effector rNTPs (185). The addition of dATP consistently inhibits activity.
Genetics and Genetic Regulation.
The E. coli RDPR is encoded by the nrd operon, a bicistronic operon consisting of nrdA and nrdB, which specify the R1 and R2 protomers, respectively. The operon was cloned, and its sequence was determined (43, 164): the order of transcription is nrdA nrdB. Mutational inactivation of protein R1 or R2 is lethal under standard aerobic growth conditions; however, both nrdA and nrdB mutants show normal growth anaerobically (18, 89, 98). Mutants with insertions in nrdB have been isolated, and these mutants not only grow anaerobically but if depleted for iron will grow under aerobic conditions as well (89).
Expression of the nrd operon is controlled at the level of transcription (85) and appears to be coupled to the initiation of DNA replication (214). Any condition that leads to a decrease in the cellular DNA/mass ratio results in increased expression of the operon, and continued protein synthesis is required in order to establish an increase in nrd mRNA during inhibition of DNA synthesis (86). The Fis protein (79) and the DnaA protein (38) bind to specific sites immediately upstream of the nrd promoter, and mutations in these regions leading to loss of binding of either protein result in a two- to threefold lowering of expression. However, the most dramatic effect on nrd expression, a five- to sixfold reduction, results from deletion of a small AT-rich region located between the Fis- and the DnaA-binding sites (15). This region is necessary for increased expression of nrd under conditions of thymine starvation (221) and is also required for cell cycle regulation of the operon (214).
The gene (nrdD) for the E. coli anaerobic rNTP reductase has been cloned, and its sequence has been determined (215). An anaerobic promoter consensus sequence, the Fnr-binding sequence (209), is located 228 bp upstream of the ATG initiation codon.
Source of Electrons for rNDP Reduction.
Electrons for reduction of rNDPs are ultimately donated by NADPH but can be shuttled to the enzyme via the small monomeric proteins thioredoxin and glutaredoxin (93). Mutants defective in thioredoxin (trxA; 139) or glutaredoxin (grx; 193) show no impairment of ribonucleotide reduction in vivo. Double mutants lacking both thioredoxin and glutaredoxin (trxA grx) maintain an active ribonucleotide reduction system (194). Thus, if thioredoxin and glutaredoxin are direct hydrogen donors for the reaction in vivo, then E. coli cells must contain at least one other hydrogen donor active with RDPR, or an alternative mechanism must exist for ribonucleotide reduction under aerobic conditions.
TSase (thyA).
Thymine nucleotides are deoxy compounds with no ribonucleotide counterparts, and they therefore cannot arise simply through the action of the ribonucleotide reductases. The homodimeric enzyme thymidylate synthase (EC 2.1.1.45; TSase) catalyzes the reductive methylation of dUMP by N 5, N 10-methylenetetrahydrofolate to yield dTMP and dihydrofolate (136). The reaction proceeds by the covalent attachment of dUMP through C-6 of the uracil moiety to a cysteine residue within the active site. Conservation of amino acid sequence is evident across numerous species and is particularly high with respect to the dUMP- and folate-binding sites (136). 5-Fluoro-dUMP is a potent inhibitor of TSase, and in the presence of the coenzyme, the enzyme binds two molecules of 5-fluoro-dUMP to form a stable, covalent ternary complex. X-ray crystallographic studies (67, 141, 142, 149) have provided detailed information on the binding of substrates and fluoro-dUMP and have led to proposals for a reaction mechanism. The E. coli thyA gene encodes a 30-kDa subunit (24); the activity of the enzyme appears to be unregulated, but studies of the production of thyA-specific transcripts indicate the potential for regulation at the level of gene expression (25). Mutants defective in TSase are obligate thymine (thymidine) auxotrophs and are resistant to the folate analogs aminopterin and trimethoprim (148).
Biosynthesis of dUMP.
E. coli and S. typhimurium can synthesize dUMP from dCDP or dUDP (148). The major pathway involves phosphorylation of dCDP to dCTP followed by deamination to dUTP and then hydrolysis to dUMP (Fig. 6). Only 25% of the cellular dUMP is derived from the phosphorylation of dUDP to dUTP and subsequent hydrolysis (Fig. 5). In accordance with functioning at a metabolic branch point, dCTP deaminase (EC 3.5.4.13; Table 5) is regulated; it shows positive homotropic cooperativity toward dCTP (the true substrate of the reaction being Mg-dCTP) and is feedback inhibited by dTTP and dUTP (21). The E. coli dcd gene encodes a polypeptide of 21.2 kDa (237), and the native enzyme from S. typhimurium has a molecular mass of 82 kDa (21), indicating that it is active as a homotetramer. The next enzyme in the pathway, dUTPase (EC 3.6.1.23; Table 5), catalyzes the hydrolysis of dUTP to dUMP and PPi. The native enzyme of E. coli is a homotrimer composed of 16-kDa subunits (43a, 203) specified by the dut gene (134). It requires Mg2+ for activity.
Both E. coli and S. typhimurium mutants lacking dCTP deaminase (dcd) have been characterized (148); their dCTP pools are increased 10-fold, and the dTTP pools are decreased by as much as 75%. Addition of thymidine or deoxyuridine restores the pools to near-normal values (168). These pool changes may explain how the cells synthesize sufficient dUMP in the absence of the contribution from dCTP deamination. A low dTTP pool will cause derepression of ribonucleotide reductase synthesis, promoting an increase in the production of dUDP. The increased dCTP pool can become a source of deoxyuridine, and hence dUMP, by hydrolysis to deoxycytidine and subsequent deamination (Fig. 4). In some strain backgrounds, growth of dcd mutants is stimulated by addition of thymidine to the medium, indicating that the cells are not capable of producing sufficient dUMP for normal growth. The acquisition of a deoA mutation suppresses this dependency on thymidine, and introduction of a cdd mutation, preventing the conversion of deoxycytidine to deoxyuridine, reestablishes it (243). The foregoing observations are consistent with the proposal discussed above for the alternative utilization of dCTP in the synthesis of dUMP.
E. coli mutants defective in dUTPase activity (dut) are impaired in the conversion of dUTP to dUMP and contain increased dUTP pools that can lead to misincorporation of uracil into DNA in place of thymine (239). Stable uracil incorporation also requires mutational inactivation of uracil-DNA N-glycosylase, the product of the ung gene. A mutant of E. coli containing a conditionally lethal dut(Ts) mutation that cannot be suppressed by the addition of thymidine or the introduction of an ung mutation has been obtained; however, suppression can be achieved by mutation of dcd (62, 237).
Conversion of dTMP to dTTP.
Two phosphorylation steps convert dTMP to dTTP. The first step is catalyzed by the highly specific dTMP kinase (Table 5; 30, 156), and the second step is catalyzed by NDK (described earlier).
Thymine and Thymidine as DNA Precursors.
Tymidine can serve as a specific DNA precursor through phosphorylation to dTMP by thymidine kinase (Table 4). Thymine may also be utilized for dTMP synthesis, since it can be converted to thymidine through condensation with deoxyribose 1-phosphate, a reaction catalyzed by thymidine phosphorylase (Table 4). Deoxyribose l-phosphate can be produced intracellularly by phosphorolytic cleavage of deoxyribonucleosides catalyzed by purine nucleoside phosphorylase (deoD) or thymidine phosphorylase (deoA) and is rapidly catabolized by the sequential action of phosphopentomutase (deoB) and deoxyriboaldolase (deoC; Fig. 4). Expression of the four deo genes that make up the deo operon is controlled by the cytR- and deoR-encoded repressors (see chapter 20 of this volume). S. typhimurium contains an inducible deoxyribokinase and a deoxyribose transport system (92), but these are apparently lacking in E. coli.
Exogenous thymine is not incorporated into DNA in thyA + cells in the absence of an endogenous deoxyribose l-phosphate pool. Addition of deoxyribonucleosides stimulates the temporary utilization of thymine by supplying deoxyribose l-phosphate. Thymidine is readily incorporated into DNA, but incorporation stops after a short time because of phosphorolysis and further catabolism of the released deoxyribose 1-phosphate. Thymidine breakdown is induced by thymidine, and consequently, high concentrations of thymidine do not promote further incorporation. Thus, deoR mutants, which contain high levels of thymidine phosphorylase and the deoxyribose l-phosphate catabolizing enzymes, cannot use thymidine as a DNA precursor. In contrast, conditions that decrease thymidine phosphorylase activity promote thymidine incorporation. These conditions include deoA mutations or the addition of uridine, a potent inhibitor of thymidine phosphorylase. Prolonged incorporation of thymidine can be achieved by using exogenous dTMP as precursor; dTMP allows for a slow feeding of thymidine, which favors phosphorylation over degradation (148).
Mutants defective in TSase (thyA) can utilize exogenous thymine but not very effectively; about 150 μM thymine is required to support normal growth. The ability of thyA mutants to utilize thymine is due to an increased production of deoxyribose l-phosphate from the increased (50- to 100-fold) dUMP pool. In thyA + cells, dUMP production is controlled by the intracellular concentration of dTTP through feedback inhibition of dCTP deaminase and regulation of ribonucleotide reductase synthesis, and therefore, cells starved for dTTP because of a defective TSase accumulate dUMP. Secondary mutations that increase the endogenous deoxyribose 1-phosphate pool in combination with an increase in the level of thymidine phosphorylase enable a thyA mutant to utilize thymine more efficiently. In contrast, mutations that lead to a reduced deoxyribose l-phosphate pool or to decreased thymidine phosphorylase activity impair thymine utilization (Table 6).
Table 6Thymine requirement in different thyA mutants |
Low concentrations of labeled thymidine of high specific activity have been used extensively as a precursor to measure short-time incorporation into nascent DNA. In interpreting such experiments, it is important to realize that artifacts springing from genetic differences among the strains used may arise. Thus, the presence of mutations affecting the uptake of thymidine (tsx, nupG, nupC) or mutations causing alterations in the internal level of thymidine phosphorylase (cytR, deoR) may give a false impression of altered DNA synthesis. A more detailed discussion of these experimental problems and how they may be overcome is given elsewhere (148).
We acknowledge the financial support of the following agencies: The Danish Center for Microbiology, the Natural Sciences and Engineering Research Council of Canada, and the NATO International Collaborative Research Program.
References
1. Abdelal, A., E. Griego, and J. L. Ingraham. 1976. Arginine-sensitive phenotype of mutations in pyrA of Salmonella typhimurium: role of ornithine carbamyltransferase in the assembly of mutant carbamylphosphate synthetase. J. Bacteriol. 128:105–113.
2. Abdelal, A., E. Griego, and J. L. Ingraham. 1978. Arginine auxotrophic phenotype of mutations in pyrA of Salmonella typhimurium: role of N-acetylornithine in the maturation of mutant carbamylphosphate synthetase. J. Bacteriol. 134:528–536.
3. Abdelal, A., and J. L. Ingraham. 1969. Cold sensitivity and other phenotypes resulting from mutation in pyrA gene. J. Biol. Chem. 244:4039–4045.
4. Abdelal, A., and J. L. Ingraham. 1975. Carbamylphosphate synthetase from Salmonella typhimurium. Regulation subunit composition, and function of the subunits. J. Biol. Chem. 250:4410–4417.
5. Aghajari, N., K. F. Jensen, and M. Gajhede. 1994. Crystallization and preliminary X-ray diffraction studies on the apo form of orotate phosphoribosyltransferase from Escherichia coli. J. Mol. Biol. 241:292–294.
6. Andersen, J. T., K. F. Jensen, and P. Poulsen. 1991. Role of transcription pausing in the control of the pyrE attenuator in Escherichia coli. Mol. Microbiol. 5:327–333.
7. Andersen, L., M. Kilstrup, and J. Neuhard. 1989. Pyrimidine, purine and nitrogen control of cytosine deaminase synthesis in Escherichia coli K12. Involvement of the glnLG and purR genes in the regulation of codA expression. Arch. Microbiol. 152:115–118.
7a. Andersen, P. S., D. Frees, R. Fast, and B. Mygind. 1995. Uracil uptake in Escherichia coli K-12: isolation of uraA mutants and cloning of the gene. J. Bacteriol. 177:2008–2013.
8. Andersen, P. S., J. M. Smith, and B. Mygind. 1992. Characterization of the upp gene encoding uracil phosphoribosyltransferase of Escherichia coli K-12. Eur. J. Biochem. 204:51–56.
9. Anderson, P. M. 1977. Binding of allosteric effectors to carbamyl-phosphate synthetase from Escherichia coli. Biochemistry 16:587–592.
10. Anderson, P. M. 1983. CTP synthetase from Escherichia coli: an improved purification procedure and characterization of hysteretic and enzyme concentration effects on kinetic properties. Biochemistry 22:3285–3292.
11. Anderson, P. M., and A. Meister. 1965. Evidence for an activated form of carbon dioxide in the reaction catalyzed by Escherichia coli carbamyl phosphate synthetase. Biochemistry 4:2803–2809.
12. Andrews, S., G. B. Cox, and F. Gibson. 1977. The anaerobic oxidation of dihydroorotate by Escherichia coli K-12. Biochim. Biophys. Acta 462:153–160.
13. Ashley, G. W., and P. A. Bartlett. 1984. Purification and properties of cytidine deaminase from Escherichia coli. J. Biol. Chem. 259:13615–13620.
14. Ashley, G. W., and P. A. Bartlett. 1984. Inhibition of Escherichia coli cytidine deaminase by a phosphapyrimidine nucleoside. J. Biol. Chem. 259:13621–13627.
15. Augustin, L. B., B. A. Jacobson, and J. A. Fuchs. 1994. Escherichia coli Fis and DnaA proteins bind specifically to the nrd promoter region and affect expression of an nrd-lac fusion. J. Bacteriol. 176:378–387.
16. Bäckström, D., R.-M. Sjöberg, and L. G. Lundberg. 1986. Nucleotide sequence of the structural gene for dihydroorotase of Escherichia coli K12. Eur. J. Biochem. 160:77–82.
17. Barilla, K. C., and R. A. Kelln. 1992. Regulation of expression of the pyrBI operon of Salmonella typhimurium LT2. Mol. Biol. (Life Sci. Adv.) 11:169–177.
18. Barlow, T. 1988. Evidence for a new ribonucleotide reductase in anaerobic E. coli. Biochem. Biophys. Res. Commun. 155:747–753.
19. Beacham, I. R., D. Haas, and E. Yagil. 1977. Mutants of Escherichia coli "cryptic" for certain periplasmic enzymes: evidence for an alteration of the outer membrane. J. Bacteriol. 129:1034–1044.
20. Beak, P., and B. Siegel. 1976. Mechanism of decarboxylation of 1,3-dimethylorotic acid. A model for orotidine 5'-monophosphate decarboxylase. J. Am. Chem. Soc. 98:3601–3606.
21. Beck, C. F., A. R. Eisenhardt, and J. Neuhard. 1975. Deoxycytidine triphosphate deaminase of Salmonella typhimurium. Purification and characterization. J. Biol. Chem. 250:609–616.
22. Beck, C. F., and J. L. Ingraham. 1971. Location on the chromosome of Salmonella typhimurium of genes governing pyrimidine metabolism. Mol. Gen. Genet. 111:303–316.
23. Beck, C. F., J. Neuhard, E. Thomassen, J. L. Ingraham, and E. Kleker. 1974. Salmonella typhimurium mutants defective in cytidine monophosphate kinase (cmk). J. Bacteriol. 120:1370–1379.
24. Belfort, M., G. F. Maley, J. Pedersen-Lane, and F. Maley. 1983. Primary structure of the Escherichia coli thyA gene and its thymidylate synthase product. Proc. Natl. Acad. Sci. USA 80:4914–4918.
25. Bell-Pedersen, D., J. L. G. Salvo, and M. Belfort. 1991. A transcription terminator in the thymidylate synthase (thyA) structural gene of Escherichia coli and construction of a viable thyA::Kmr deletion. J. Bacteriol. 173:1193–1200.
26. Betts, L., S. Xiang, S. A. Short, R. Wolfenden, and C. C. Carter. 1994. Cytidine deaminase: 2.3 Å crystal structure of an enzyme:transition state analog complex. J. Mol. Biol. 235:635–656.
27. Bhatia, M. B., and C. Grubmeyer. 1993. The role of divalent magnesium in activating the reaction catalyzed by orotate phosphoribosyltransferase. Arch. Biochem. Biophys. 303:321–325.
28. Bhatia, M. B., A. Vinitsky, and C. Grubmeyer. 1990. Kinetic mechanism of orotate phosphoribosyltransferase from Salmonella typhimurium. Biochemistry 29:10480–10487.
29. Bianchi, V., P. Reichard, R. Eliasson, E. Pontis, M. Krook, H. Jörnvall, and E. Haggård-Ljungquist. 1993. Escherichia coli ferredoxin NADP reductase: activation of E. coli anaerobic ribonucleotide reduction, cloning of the gene (fpr), and overproduction of the protein. J. Bacteriol. 175:1590–1595.
30. Binkley, J. P., and P. L. Kuempel. 1986. Genetic mapping in Escherichia coli of tmk, the locus for dTMP kinase. J. Bacteriol. 168:1457–1458.
31. Black, M. E., and D. E. Hruby. 1991. Nucleotide sequence of the Escherichia coli thymidine kinase gene provides evidence for conservation of functional domains and quaternary structure. Mol. Microbiol. 5:373–379.
32. Boettcher, B., and A. Meister. 1982. Regulation of Escherichia coli carbamyl phosphate synthetase. Evidence for overlap of the allosteric nucleotide binding sites. J. Biol. Chem. 257:13971–13976.
33. Boettcher, B. R., and A. Meister. 1980. Covalent modification of the active site of carbamyl phosphate synthetase by 5'-p-fluorosulfonylbenzoyladenosine. Direct evidence for two functionally different ATP-binding sites. J. Biol. Chem. 255:7129–7133.
34. Bonekamp, F., H. D. Andersen, T. Christensen, and K. F. Jensen. 1985. Codon-defined ribosomal pausing in Escherichia coli detected by using the pyrE attenuator to probe the coupling between transcription and translation. Nucleic Acids Res. 13:4113–4123.
35. Bonekamp, F., K. Clemmesen, O. Karlström, and K. F. Jensen. 1984. Mechanism of UTP-modulated attenuation at the pyrE gene of Escherichia coli: an example of operon polarity control through the coupling of translation to transcription. EMBO J. 3:2857–2861.
36. Bothwell, M., and H. K. Schachman. 1980. A model for the assembly of aspartate transcarbamoylase from catalytic and regulatory subunits. J. Biol. Chem. 255:1971–1977.
37. Bouvier, J., J.-C. Patte, and P. Stragier. 1984. Multiple regulatory signals in the control region of the Escherichia coli carAB operon. Proc. Natl. Acad. Sci. USA 81:4139–4143.
38. Bramhill, D., and A. Kornberg. 1988. A model for initiation at origins of DNA replication. Cell 54:915–918.
39. Bremer, E., A. Middendorf, J. Matinussen, and P. Valentin-Hansen. 1990. Analysis of the tsx gene, which encodes a nucleoside-specific channel-forming protein (Tsx) in the outer membrane of Escherichia coli. Gene 96:59–65.
40. Brown, D. C., and K. D. Collins. 1991. Dihydroorotase from Escherichia coli. Substitution of Co(II) for the active site Zn(II). J. Biol. Chem. 266:1597–1604.
41. Burns, D. L., and H. K. Schachman. 1982. Ligand-promoted strengthening of interchain bonding domains in catalytic subunits of aspartate transcarbamoylase. J. Biol. Chem. 257:12214–12218.
42. Bussey, L. B., and J. L. Ingraham. 1982. A regulatory gene (use) affecting the expression of pyrA and certain other pyrimidine genes. J. Bacteriol. 151:144–152.
43. Carlson, J., J. A. Fuchs, and J. Messing. 1984. Primary structure of the Escherichia coli ribonucleoside diphosphate reductase operon. Proc. Natl. Acad. Sci. USA 81:4294–4297.
43a. Cedergren-Zeppezauer, E. S., G. Larsson, P. P. Nyman, Z. Dauter, and K. S. Wilson. 1992. Crystal structure of a dUTPase. Nature (London) 355:740–743.
44. Charlier, D., N. Huysveld, M. Roovers, and N. Glansdorff. 1994. On the role of the Escherichia coli integration host factor (IHF) in repression at a distance of the pyrimidine specific promoter P1 of the carAB operon. Biochimie 76:1041–1051.
45. Charlier, D., M. Roovers, D. Gigot, N. Huysveld, A. Pierard, and N. Glansdorff. 1993. Integration host factor (IHF) modulates the expression of the pyrimidine-specific promoter of the carAB operons of Escherichia coli K12 and Salmonella typhimurium LT2. Mol. Gen. Genet. 237:273–286.
46. Charlier, D., M. Roovers, D. Gigot, J. Van Handenhove, A. Pierard, and N. Glansdorff. 1992. Arginine regulon of Escherichia coli K-12: a study of repressor-operator interactions and of in vitro binding affinities versus in vivo repression. J. Mol. Biol. 226:367–386.
47. Charlier, D., G. Weyens, M. Roovers, J. Piette, C. Bocquet, A. Pierard, and N. Glansdorff. 1988. Molecular interactions in the control region of the carAB operon encoding Escherichia coli carbamoylphosphate synthetase. J. Mol. Biol. 204:867–877.
48. Chen, M. S., and W. H. Prusoff. 1978. Thymidine kinase from Escherichia coli. Methods Enzymol. 51:354–360.
49. Choi, K. Y., and H. Zalkin. 1990. Regulation of Escherichia coli pyrC by the purine regulon repressor protein. J. Bacteriol. 172:3201–3207.
50. Clemmesen, K., F. Bonekamp, O. Karlström, and K. F. Jensen. 1985. Role of translation in the UTP-modulated attenuation at the pyrBI operon of Escherichia coli. Mol. Gen. Genet. 201:247–251.
51. Cohen, R. M., and R. Wolfenden. 1971. Cytidine deaminase from Escherichia coli: purification, properties, and inhibition by the potential transition state analog 3,4,5,6-tetrahydrouridine. J. Biol. Chem. 246:7561–7565.
52. Cook, W. J., G. W. Kozalka, W. W. Hall, S. V. L. Narayana, and S. E. Ealick. 1987. Crystallization and preliminary X-ray investigation of uridine phosphorylase from Escherichia coli. J. Biol. Chem. 262:2852–2853.
53. Corder, T. S., and J. R. Wild. 1989. Discrimination between nucleotide effector responses of aspartate transcarbamoylase due to a single site substitution in the allosteric binding site. J. Biol. Chem. 264:7425–7430.
53a. Craig, J. E., Y. Zhang, and M.P. Gallagher. 1994. Cloning of the nupC gene of Escherichia coli encoding a nucleoside transport system, and identification of an adjacent insertion element, IS 186. Mol. Microbiol. 11:1159–1168.
54. Cunin, R., N. Glansdorff, A. Pierard, and V. Stalon. 1986. Arginine biosynthesis and metabolism in bacteria. Microbiol. Rev. 50:314–352.
55. Dandanell, G., K. Norris, and K. Hammer. 1991. Long-distance deoR regulation of gene expression in Escherichia coli. Ann. N.Y. Acad. Sci. 646:19–30.
56. Danielsen, S., M. Kilstrup, K. Barilla, B. Jochimsen, and J. Neuhard. 1992. Characterization of the Escherichia coli codBA operon encoding cytosine permease and cytosine deaminase. Mol. Microbiol. 6:1335–1344.
57. Donahue, J. P., and C. L. Turnbough. 1990. Characterization of transcriptional initiation from promoters P1 and P2 of the pyrBI operon of Escherichia coli K-12. J. Biol. Chem. 265:19091–19099.
58. Donovan, W. P., and S. R. Kushner. 1983. Cloning and physical analysis of the pyrF gene (coding for orotidine-5'-phosphate decarboxylase) from Escherichia coli K-12. Gene 25:39–48.
59. Donovan, W. P., and S. R. Kushner. 1983. Purification and characterization of orotidine-5'-phosphate decarboxylase from Escherichia coli K-12. J. Bacteriol. 156:620–624.
60. Drabikowska, A. K., and G. Wozniak. 1990. Modification of uridine phosphorylase from Escherichia coli by diethyl pyrocarbonate. Biochem. J. 270:319–323.
61. Eisenstein, E., M. S. Han, T. S. Woo, J. M. Ritchy, I. Gibbons, Y. R. Yang, and H. K. Schachman. 1992. Negative complementation in aspartate transcarbamylase. Analysis of hybrid enzyme molecules containing different arrangements of polypeptide chains from wild-type and inactive mutant catalytic subunits. J. Biol. Chem. 267:22148–22155.
62. El-Hajj, H. H., L. Wang, and B. Weiss. 1992. Multiple mutant Escherichia coli synthesizing virtually thymineless DNA during limited growth. J. Bacteriol. 174:4450–4456.
63. Eliasson, R., E. Pontis, M. Fontecave, C. Gerez, J. Harder, H. Jörnvall, M. Krook, and P. Reichard. 1992. Characterization of components of the anaerobic ribonucleotide reductase system from Escherichia coli. J. Biol. Chem. 267:25541–25547.
64. England, P., and G. Hervé. 1992. Synergistic inhibition of Escherichia coli aspartate transcarbamylase by CTP and UTP: binding studies using continuous-flow dialysis. Biochemistry 31:9725–9732.
65. England, P., and G. Hervé. 1994. Involvement of the γ-phosphate of UTP in the synergistic inhibition of Escherichia coli aspartate transcarbamylase by CTP and UTP. Biochemistry 33:3913–3918.
66. Fast, R., and O. Skjöld. 1977. Biochemical mechanism of uracil uptake regulation in Escherichia coli B. Allosteric effects on uracil phosphoribosyltransferase under stringent conditions. J. Biol. Chem. 252:7620–7624.
67. Finer-Moore, J. S., W. R. Montfort, and R. M. Stroud. 1990. Pairwise specificity and sequential binding in enzyme catalysis: thymidylate synthase. Biochemistry 29:6977–6986.
68. Fontecave, M., P. Nordlund, H. Eklund, and P. Reichard. 1992. The redox centers of ribonucleotide reductase of Escherichia coli. Adv. Enzymol. 65:147–183.
69. Frick, L., C. Yang, V. E. Marquez, and R. Wolfenden 1989. Binding of pyrimidin-2-one ribonucleoside by cytidine deaminase as the transition-state analogue 3,4-dihydrouridine and the contribution of the 4-hydroxyl group to its binding affinity. Biochemistry 28:9423–9430.
70. Frick, M. M., J. Neuhard, and R. A. Kelln. 1990. Cloning, nucleotide sequence and regulation of the Salmonella typhimurium pyrD gene encoding dihydroorotate dehydrogenase. Eur. J. Biochem. 194:573–578.
70a. Fricke, J., J. Neuhard, R. A. Kelln, and S. Pedersen. 1995. The cmk gene encoding cytidine monophosphate kinase is located in the rpsA operon and is required for normal replication rate in Escherichia coli. J. Bacteriol. 177:517–523.
71. Gerhart, J. C. 1970. A discussion of the regulatory properties of aspartate transcarbamylase from Escherichia coli. Curr. Top. Cell. Regul. 2:275–325.
72. Gerlach, P., L. Søgaard-Andersen, H. Pedersen, J. Martinussen, P. Valentin-Hansen, and E. Bremer. 1991. The cyclic AMP (cAMP)-cAMP receptor protein complex functions both as an activator and as a corepressor at the tsx-p 2 promoter of Escherichia coli K-12. J. Bacteriol. 173:5419–5430.
73. Ginther, C. L., and J. L. Ingraham. 1974. Nucleoside diphosphokinase of Salmonella typhimurium. J. Biol. Chem. 249:3406–3411.
74. Gouaux, J. E., K. L. Krause, and W. N. Lipscomb. 1987. The catalytic mechanism of Escherichia coli aspartate carbamoyltransferase: a molecular modelling study. Biochem. Biophys. Res. Commun. 142:893–897.
75. Gouaux, J. E., and W. N. Lipscomb. 1990. Crystal structures of phosphonoacetamide ligated T and phosphonoacetamide and malonate ligated R states of aspartate carbamoyltransferase at 2.8-Å resolution and neutral pH. Biochemistry 29:389–402.
76. Gouaux, J. E., R. C. Stevens, and W. N. Lipscomb. 1990. Crystal structures of aspartate carbamoyltransferase ligated with phosphonoacetamide, malonate, and CTP or ATP at a 2.8 Å resolution and neutral pH. Biochemistry 29:7702–7715.
77. Grubmeyer, C., E. Segura, and R. Dorfman. 1993. Active site lysines in orotate phsophoribosyltransferase. J. Biol. Chem. 268:20299–20304.
78. Guillou, F., S. D. Rubino, R. S. Markovitz, D. M. Kinney, and C. J. Lusty. 1989. Escherichia coli carbamoyl-phosphate synthetase: domains of glutaminase and synthetase subunit interaction. Proc. Natl. Acad. Sci. USA 86:8304–8308.
79. Haffter, P., and T. A. Bickle. 1987. Purification and DNA-binding properties of FIS and CIN, two proteins required for bacteriophage P1 site specific recombination system, cin. J. Mol. Biol. 198:579–587.
80. Hama, H., N. Almaula, C. G. Lerner, S. Inouye, and M. Inouye. 1991. Nucleoside diphosphate kinase from Escherichia coli; its overproduction and sequence comparison with eukaryotic enzymes. Gene 105:31–36.
81. Hammer-Jespersen, K. 1983. Nucleoside catabolism, p. 203–258. In A. Munch-Petersen (ed.), Metabolism of Nucleotides, Nucleosides and Nucleobases in Microorganisms. Academic Press, Inc. (London), Ltd., London.
82. Hammer-Jespersen, K., and A. Munch-Petersen. 1973. Mutants of Escherichia coli unable to metabolize cytidine: isolation and characterization. Mol. Gen. Genet. 126:177–186.
83. Han, B.-D., W. G. Nolan, H. P. Hopkins, R. T. Jones, J. L. Ingraham, and A. T. Abdelal. 1990. Effect of growth temperature on folding of carbamoylphosphate synthetase of Salmonella typhimurium and a cold-sensitive derivative. J. Bacteriol. 172:5089–5096.
84. Handschumacher, R. E. 1960. Orotidylic acid decarboxylase: inhibition studies with azauridine 5'-phosphate. J. Biol. Chem. 235:2917–2919.
85. Hanke, P. D., and J. A. Fuchs. 1983. Regulation of ribonucleoside diphosphate reductase mRNA synthesis in Escherichia coli. J. Bacteriol. 154:1040–1045.
86. Hanke, P. D., and J. A. Fuchs. 1984. Requirement of protein synthesis for the induction of ribonucleoside diphosphate reductase mRNA in Escherichia coli. Mol. Gen. Genet. 193:327–331.
87. Hansen, S. E. W., N. Jensen, and A. Munch-Petersen. 1987. Studies on the sequence and structure of the Escherichia coli K-12 nupG gene, encoding a nucleoside-transport system. Eur. J. Biochem. 168:385–391.
88. Hantke, K. 1976. Phage T6-colicin K receptor and nucleoside transport in Escherichia coli. FEBS Lett. 70:109–112.
89. Hantke, K. 1988. Characterization of an iron sensitive Mud1 mutant in E. coli lacking the ribonucleotide reductase subunit B2. Arch. Microbiol. 149:344–349.
90. Harder, J., R. Eliasson, E. Pontis, M. D. Ballinger, and P. Reichard. 1992. Activation of the anaerobic ribonucleotide reductase from Escherichia coli by S-adenosylmethionine. J. Biol. Chem. 267:25548–25552.
91. Hervé, G. 1989. Aspartate transcarbamylase from Escherichia coli, p. 61–79. In G. Hervé (ed.), Allosteric Enzymes. CRC Press, Inc., Boca Raton, Fla.
92. Hoffee, P. A. 1968. 2-Deoxyribose gene-enzyme complex in Salmonella typhimurium. I. Isolation and enzymatic characterization of 2-deoxyribose-negative mutants. J. Bacteriol. 95:449–457.
93. Holmgren, A. 1989. Thioredoxin and glutaredoxin systems. J. Biol. Chem. 264:13963–13966.
94. Hosono, H., and S. Kuno. 1973. The purification and properties of cytidine deaminase from Escherichia coli. J. Biochem. 74:797–803.
95. Hove-Jensen, B., K. W. Harlow, C. J. King, and R. L. Switzer. 1986. Phosphoribosylpyrophosphate synthetase of Escherichia coli. Properties of the purified enzyme and primary structure of the prs gene. J. Biol. Chem. 261:6765–6771.
96. Hsuanyu, Y., F. C. Wedler, E. R. Kantrowitz, and S. A. Middleton. 1989. Site-specific mutation of Tyr240-Phe in the catalytic chain of Escherichia coli aspartate transcarbamylase. Consequences for kinetic mechanism. J. Biol. Chem. 264:17259–17265.
97. Ingraham, J. L., and J. Neuhard. 1972. Cold-sensitive mutants of Salmonella typhimurium defective in uridine monophosphate kinase (pyrH). J. Biol. Chem. 247:6259–6265.
98. Jamison, C. S., and H. I. Adler. 1987. Mutations in Escherichia coli that effect sensitivity to oxygen. J. Bacteriol. 169:5087–5094.
99. Jensen, K. F. 1979. Apparent involvement of purines in the control of expression of Salmonella typhimurium pyr genes: analysis of a leaky guaB mutant resistant to pyrimidine analogs. J. Bacteriol. 138:731–738.
100. Jensen, K. F. 1988. Hyper-regulation of pyr gene expression in Escherichia coli with slow ribosomes. Evidence for RNA polymerase pausing in vivo. Eur. J. Biochem. 175:587–593.
101. Jensen, K. F. 1989. Regulation of Salmonella typhimurium pyr gene expression: effect of changing both purine and pyrimidine nucleotide pools. J. Gen. Microbiol. 135:805–815.
102. Jensen, K. F. 1993. The Escherichia coli "wild types" W3110 and MG1655 have an rph frameshift mutation that leads to pyrimidine starvation due to low pyrE expression levels. J. Bacteriol. 175:3401–3407.
103. Jensen, K. F., R. Fast, O. Karlström, and J. N. Larsen. 1986. Association of RNA polymerase having increased Km for ATP and UTP with hyperexpression of the pyrB and pyrE genes of Salmonella typhimurium. J. Bacteriol. 166:857–865.
104. Jensen, K. F., J. N. Larsen, L. Schack, and A. Sivertsen. 1984. Studies on the structure and expression of Escherichia coli pyrC, pyrD, and pyrF using the cloned genes. Eur. J. Biochem. 140:343–352.
105. Jensen, K. F., J. Neuhard, and L. Schack. 1982. RNA polymerase involvement in the regulation of expression of Salmonella typhimurium pyr genes. Isolation and characterization of a fluorouracil-resistant mutant with high, constitutive expression of the pyrB and pyrE genes due to a mutation in rpoBC. EMBO J. 1:69–74.
105a. Jordan, A., I. Gibert, and J. Barbe. 1994. Cloning and sequencing of the genes from Salmonella typhimurium encoding a new bacterial ribonucleotide reductase. J. Bacteriol. 176:3420–3427.
106. Justesen, J., and J. Neuhard. 1975. pyrR identical to pyrH in Salmonella typhimurium: control of expression of the pyr genes. J. Bacteriol. 123:851–854.
107. Kantrowitz, E. R., and W. N. Lipscomb. 1990. Escherichia coli aspartate transcarbamoylase: the molecular basis for a concerted allosteric transition. Trends Biochem. Sci. 15:53–59.
108. Kantrowitz, E. R., S. C. Pastra-Landis, and W. N. Lipscomb. 1980. E. coli aspartate transcarbamylase. Part I. Catalytic and regulatory functions. Trends Biochem. Sci. 5:124–128.
109. Kantrowitz, E. R., S. C. Pastra-Landis, and W. N. Lipscomb. 1980. E. coli aspartate transcarbamylase. Part II. Structure and allosteric interactions. Trends Biochem. Sci. 5:150–153.
110. Karibian, D. 1978. Dihydroorotate dehydrogenase (Escherichia coli). Methods Enzymol. 51:58–63.
111. Karibian, D., and P. Couchoud. 1974. Dihydro-orotate oxidase of Escherichia coli K12: purification, properties, and relation to the cytoplasmic membrane. Biochim. Biophys. Acta 364:218–232.
112. Katsuragi, T., T. Sakai, K. Matsumoto, and K. Tonomura. 1986. Cytosine deaminase from Escherichia coli—production, purification, and some characteristics. Agric. Biol. Chem. 50:1721–1730.
113. Ke, H., W. N. Lipscomb, Y. Cho, and R. B. Honzatko. 1988. Complex of N-phosphonoacetyl-l-aspartate with aspartate carbamoyltransferase. J. Mol. Biol. 204:725–747.
114. Kelln, R. A., J. J. Kinahan, K. F. Foltermann, and G. A. O’Donovan. 1975. Pyrimidine biosynthetic enzymes of Salmonella typhimurium, repressed specifically by growth in the presence of cytidine. J. Bacteriol. 124:764–774.
115. Kelln, R. A., and J. Neuhard. 1988. Regulation of pyrC expression in Salmonella typhimurium: identification of a regulatory region. Mol. Gen. Genet. 212:287–294.
116. Kerr, C. T., and R. W. Miller. 1968. Dihydroorotate-ubiquinone reductase complex of Escherichia coli B. J. Biol. Chem. 243:2963–2968.
117. Kilstrup, M., C.-D. Lu, A. Abdelal, and J. Neuhard. 1988. Nucleotide sequence of the carA gene and regulation of the carAB operon in Salmonella typhimurium. Eur. J. Biochem. 176:421–429.
118. Kilstrup, M., L. M. Meng, J. Neuhard, and P. Nygaard. 1989. Genetic evidence for a repressor of synthesis of cytosine deaminase and purine biosynthesis enzymes in Escherichia coli. J. Bacteriol. 171:2124–2127.
119. Kimsey, H. H., and D. Kaiser. 1992. The orotidine-5'-monophosphate decarboxylase gene of Myxococcus xanthus. Comparison of the OMP decarboxylase gene family. J. Biol. Chem. 267:819–824.
120. Kleanthous, C., D. E. Wemmer, and H. K. Schachman. 1988. The role of an active site histidine in the catalytic mechanism of aspartate transcarbamoylase. J. Biol. Chem. 263:13062–13067.
121. Koshland, D. E., and A. Levitzki. 1974. CTP synthetase and related enzymes, p. 539–559. In P. D. Boyer (ed.), The Enzymes, vol. 10. Academic Press, Inc., New York.
122. Krause, K. L., K. W. Voltz, and W. N. Lipscomb. 1987. 2.5 Å structure of aspartate carbamoyltransferase complexed with the bisubstrate analogue N-(phosphonoacetyl)-l-aspartate. J. Mol. Biol. 193:527–553.
123. Larsen, J. N., and K. F. Jensen. 1985. Nucleotide sequence of the pyrD gene of Escherichia coli and characterization of the flavoprotein dihydroorotate dehydrogenase. Eur. J. Biochem. 151:59–65.
124. Larsson, Å., and B.-M. Sjöberg. 1986. Identification of the stable free radical tyrosine residue in ribonucleotide reductase. EMBO J. 5:2037–2040.
125. Leer, J. C., K. Hammer-Jespersen, and M. Schwartz. 1977. Uridine phosphorylase from Escherichia coli. Physical and chemical characterization. Eur. J. Biochem. 75:217–224.
126. Legrain, C., V. Stalon, N. Glansdorff, D. Gigot, A. Pierard, and M. Crabeel. 1976. Structural and regulatory mutations allowing utilization of citrulline or carbamoylaspartate as a source of carbamoylphosphate in Escherichia coli K-12. J. Bacteriol. 128:39 48.
127. Levin, H. L., K. Park, and H. K. Schachman. 1989. Attenuation in the regulation of the pyrBI operon of Escherichia coli: in vivo studies of transcriptional termination. J. Biol. Chem. 264:14638–14645.
128. Levin, H. L., and H. K. Schachman. 1985. Regulation of aspartate transcarbamoylase synthesis in Escherichia coli: analysis of deletion mutations in the promoter region of the pyrBI operon. Proc. Natl. Acad. Sci. USA 82:4643–4647.
129. Lichtenstein, J., H. D. Barner, and S. S. Cohen. 1960. The metabolism of exogenously supplied nucleotides by Escherichia coli. J. Biol. Chem. 235:457–465.
130. Liu, C., and C. L. Turnbough. 1989. Multiple control mechanisms for pyrimidine-mediated regulation of pyrBI operon expression in Escherichia coli K-12. J. Bacteriol. 171:3337–3342.
131. Lu, C.-D., and A. T. Abdelal. 1993. The Salmonella typhimurium uracil-sensitive mutation use is in argU and encodes a minor arginine tRNA. J. Bacteriol. 175:3897–3899.
132. Lu, C.-D., J. E. Houghton, and A. T. Abdelal. 1992. Characterization of the arginine repressor of Salmonella typhimurium and its interaction with the carAB operator. J. Mol. Biol. 225:11–24.
133. Lu, C.-D., M. Kilstrup, J. Neuhard, and A. Abdelal. 1989. Pyrimidine regulation of tandem promoters for carAB in Salmonella typhimurium. J. Bacteriol. 171:5436–5442.
134. Lundberg, L. G., H.-O. Thoresson, O. H. Karlström, and P.-O. Nyman. 1983. Nucleotide sequence of the structural gene for dUTPase of Escherichia coli K-12. EMBO J. 2:967–971.
135. Machida, H., and A. Kuninaka. 1969. Studies on the accumulation of orotic acid by Escherichia coli K12. Agric. Biol. Chem. 33:868–875.
136. Maley, F., and G. F. Maley. 1990. A tale of two enzymes, deoxycytidylate deaminase and thymidylate synthase. Prog. Nucleic Acid Res. Mol. Biol. 39:49–80.
137. Mao, S. S., T. P. Holler, G. X. Yu, J. M. Bollinger, S. Booker, M. I. Johnston, and J. Stubbe. 1992. A model for the role of multiple cysteine residues involved in ribonucleotide reduction: amazing and still confusing. Biochemistry 31:9733–9743.
138. Mao, S. S., G. X. Yu, D. Chalfoun, and J. Stubbe. 1992. Characterization of C439CR1, a mutant of Escherichia coli ribonucleotide diphosphate reductase: evidence that C439 is a residue essential for nucleotide reduction and C439SR1 is a protein possessing novel thioredoxin-like activity. Biochemistry 31:9752–9759.
139. Mark, D. F., J. W. Chase, and C. C. Richardson. 1977. Genetic mapping of trxA, a gene affecting thioredoxin in Escherichia coli K12. Mol. Gen. Genet. 155:145–152.
140. Markby, D. W., B. B. Zhou, and H. K. Schachman. 1991. A 70-amino acid zinc-binding polypeptide from the regulatory chain of aspartate transcarbamoylase forms a stable complex with the catalytic subunit leading to markedly altered enzyme activity. Proc. Natl. Acad. Sci. USA 88:10568–10572.
141. Matthews, D. A., K. Appelt, S. J. Oatley, and N. H. Xuong. 1990. Crystal structure of Escherichia coli thymidylate synthase containing bound 5-fluoro-2'-deoxyuridylate and 10-propargyl-5,8-dideazafolate. J. Mol. Biol. 214:923–936.
142. Matthews, D. A., J. E. Villafranca, C. A. Janson, W. W. Smith, K. Welsh, and S. Freer. 1990. Stereochemical mechanism of action for thymidylate synthase based on the X-ray structure of the covalent inhibitory ternary complex with 5-fluoro-2'deoxyuridylate and 5,10-methylenetetrahydrofolate. J. Mol. Biol. 214:937–948.
143. McCarthy, M. P., and N. M. Allewell. 1983. Thermodynamics of assembly of E. coli aspartate transcarbamoylase. Proc. Natl. Acad. Sci. USA 80:6824–6828.
144. McKeown, M., M. Kahn, and P. Hanawalt. 1976. Thymidine uptake and utilization in Escherichia coli: a new gene controlling nucleoside transport. J. Bacteriol. 126:814–822.
145. Meister, A. 1989. Mechanism and regulation of the glutamine-dependent carbamyl phosphate synthetase of Escherichia coli. Adv. Enzymol. 62:315–374.
146. Mergeay, M., D. Gigot, J. Beckmann, N. Glansdorff, and A. Pierard. 1974. Physiology and genetics of carbamoylphosphate synthesis in Escherichia coli K12. Mol. Gen. Genet. 133:299–316.
147. Michaels, G., R. A. Kelln, and F. E. Nargang. 1987. Cloning, nucleotide sequence and expression of the pyrBI operon of Salmonella typhimurium. Eur. J. Biochem. 166:55–61.
148. Møllgaard, H., and J. Neuhard. 1983. Biosynthesis of deoxythymidine triphosphate, p. 149–198. In A. Munch-Petersen (ed.), Metabolism of Nucleotides, Nucleosides, and Nucleobases in Microorganisms. Academic Press, Inc. (London), Ltd., London.
149. Montfort, W. R., K. M. Perry, E. B. Fauman, J. S. Finer-Moore, G. F. Maley, L. Hardy, F. Maley, and R. M. Stroud. 1990. Structure, multiple site binding, and segmental accommodation in thymidylate synthase on binding dUMP and an anti-folate. Biochemistry 29:6964–6977.
150. Mulliez, E., M. Fontecave, J. Gaillard, and P. Reichard. 1993. An iron-sulfur center and a free radical in the active anaerobic ribonucleotide reductase of Escherichia coli. J. Biol. Chem. 268:2296–2299.
151. Munch-Petersen, A., and N. Jensen. 1990. Analysis of the regulatory region of the Escherichia coli nupG gene, encoding a nucleoside-transport protein. Eur. J. Biochem. 190:547–551.
152. Munch-Petersen, A., and B. Mygind. 1983. Transport of nucleic acid precursors, p. 259–305. In A. Munch-Petersen (ed.), Metabolism of Nucleotides, Nucleosides, and Nucleobases in Microorganisms. Academic Press, Inc. (London), Ltd., London.
153. Nagy, M., F. Lacroute, and D. Thomas. 1992. Divergent evolution of pyrimidine biosynthesis between anaerobic and aerobic yeasts. Proc. Natl. Acad. Sci. USA 89:8966–8970.
154. Navre, M., and H. K. Schachman. 1983. Synthesis of aspartate transcarbamoylase in Escherichia coli: transcriptional regulation of the pyrB-pyrI operon. Proc. Natl. Acad. Sci. USA 80:1207–1211.
155. Nelbach, M. E., V. P. Pigiet, J. C. Gerhart, and H. K. Schachman. 1972. A role for zinc in the quaternary structure of aspartate transcarbamylase from Escherichia coli. Biochemistry 11:315–327.
156. Nelson, D. P., and C. E. Carter. 1969. Purification and characterization of thymidine 5'-monophosphate kinase from Escherichia coli B. J. Biol. Chem. 244:5254–5262.
157. Neuhard, J. 1983. Utilization of preformed pyrimidine bases and nucleosides, p. 95–148. In A. Munch-Petersen (ed.), Metabolism of Nucleotides, Nucleosides, and Nucleobases in Microorganisms. Academic Press, Inc. (London), Ltd., London.
158. Neuhard J., K. F. Jensen, and E. Stauning. 1982. Salmonella typhimurium mutants with altered expression of the pyrA gene due to changes in RNA polymerase. EMBO J. 9:1141–1145.
159. Neuhard, J., and R. A. Kelln. 1988. A chromosomal mutation mediating increased expression of pyrE in Salmonella typhimurium is located within the proposed attenuator. Can. J. Microbiol. 34:686–687.
160. Neuhard, J., R. A. Kelln, and E. Stauning. 1986. Cloning and structural characterization of the Salmonella typhimurium pyrC gene encoding dihydroorotase. Eur. J. Biochem. 157:335–342.
161. Neuhard, J., L. M. Meng, K. I. Sørensen, and R. A. Kelln. 1990. Purine control of pyrC expression in Salmonella typhimurium. Int. J. Purine Pyrimidine Res. 1:61–66.
162. Neuhard, J., E. Stauning and R. A. Kelln. 1985. Cloning and characterization of the pyrE gene and of pyrE::Mud1(ApR lac) fusions from Salmonella typhimurium. Eur. J. Biochem. 146:597–603.
163. Newton, C. J., and E. R. Kantrowitz. 1990. Importance of domain closure for homotropic cooperativity in Escherichia coli aspartate transcarbamylase. Biochemistry 29:1444–1451.
164. Nilsson, O., A. Åberg, T. Lundquist, and B.-M. Sjöberg. 1988. Nucleotide sequence of the gene coding for the large subunit of ribonucleotide reductase of Escherichia coli. Nucleic Acids Res. 16:4174. (Correction.)
165. Nordlund, P., and H. Eklund. 1994. Structure and function of the Escherichia coli ribonucleotide reductase protein R2. J. Mol. Biol. 232:123–164.
166. Nygaard, P. 1973. Nucleoside-catabolizing enzymes in Salmonella typhimurium. Induction by ribonucleosides. Eur. J. Biochem. 36:267–272.
167. Nyunoya, H., and C. J. Lusty. 1983. The carB gene of Escherichia coli: a duplicated gene coding for the large subunit of carbamoyl-phosphate synthetase. Proc. Natl. Acad. Sci. USA 80:4629–4633.
168. O’Donovan, G. A., G. Edlin, J. A. Fuchs, J. Neuhard, and E. Thomassen. 1971. Deoxycytidine triphosphate deaminase: characterization of an Escherichia coli mutant deficient in the enzyme. J. Bacteriol. 105:666–672.
169. O’Donovan, G. A., H. Holoubek, and J. C. Gerhart. 1972. Regulatory properties of intergeneric hybrids of aspartate transcarbamylase. Nature (London) New Biol. 238:264–266.
170. Ohtsuki, K., M. Yokoyama, T. Koike, and N. Ishida. 1984. Nucleoside diphosphate kinase in Escherichia coli: its polypeptide structure and reaction intermediate. Biochem. Int. 8:715–723.
171. Ost, K. A., and M. P. Deutscher. 1991. Escherichia coli orfE (upstream of pyrE) encodes RNase PH. J. Bacteriol. 173:5589–5591.
172. Peterson, C. B., and H. K. Schachman. 1992. Long range effects of amino acid substitutions in the catalytic chain of aspartate transcarbamoylase. Localized replacements in the carboxy-terminal α-helix cause marked alterations in allosteric properties and intersubunit interactions. J. Biol. Chem. 267:2443–2450.
173. Pierard, A., N. Glansdorff, D. Gigot, M. Crabeel, P. Halleux, and L. Thiry. 1976. Repression of Escherichia coli carbamoylphosphate synthase: relationships with enzyme synthesis in the arginine and pyrimidine pathways. J. Bacteriol. 127:291–301.
174. Pierard, A., N. Glansdorff, M. Mergeay, and J. M. Wiame. 1965. Control of the biosynthesis of carbamoyl phosphate in Escherichia coli. J. Mol. Biol. 14:23–36.
175. Pierard, A., N. Glansdorff, and J. Yashphe. 1972. Mutations affecting uridine monophosphate pyrophosphorylase or the argR gene in Escherichia coli. Effects on carbamoyl phosphate and pyrimidine biosynthesis and on uracil uptake. Mol. Gen. Genet. 118:235–245.
176. Piette, J., H. Nyunoya, C. J. Lusty, R. Cunin, G. Weyens, M. Crabeel, D. Charlier, N. Glansdorff, and A. Pierard. 1984. DNA sequence of the carA gene and the control region of carAB: tandem promoters, respectively controlled by arginine and the pyrimidines, regulate the synthesis of carbamoyl-phosphate synthetase in Escherichia coli K-12. Proc. Natl. Acad. Sci. USA 81:4134 4138.
177. Porter, D. J. T., and E. A. Austin. 1993. Cytosine deaminase. The roles of divalent metal ions in catalysis. J. Biol. Chem. 268:24005–24011.
178. Post, L.E., D. J. Post, and F. M. Raushel. 1990. Dissection of the functional domains of Escherichia coli carbamoyl phosphate synthetase by site-directed mutagenesis. J. Biol. Chem. 265:7742–7747.
179. Poulsen, P., F. Bonekamp, and K. F. Jensen. 1984. Structure of the Escherichia coli pyrE operon and control of pyrE expression by a UTP modulated intercistronic attenuation. EMBO J. 3:1783–1790.
180. Poulsen, P., and K. F. Jensen. 1987. Effect of UTP and GTP pools on attenuation at the pyrE gene of Escherichia coli. Mol. Gen. Genet. 208:152–158.
181. Poulsen, P., K. F. Jensen, P. Valentin-Hansen, P. Carlsson, and L. G. Lundberg. 1983. Nucleotide sequence of thc Escherichia coli pyrE gene and of the DNA in front of the protein-coding region. Eur. J. Biochem. 135:223–229.
182. Quinn, C. L., B. T. Stephenson, and R. L. Switzer. 1991. Functional organization and nucleotide sequence of the Bacillus subtilis pyrimidine biosynthetic operon. J. Biol. Chem. 266:9113–9127.
183. Rasmussen, U. B., B. Mygind, and P. Nygaard. 1986. Purification and some properties of uracil phosphoribosyltransferase from Escherichia coli K-12. Biochim. Biophys. Acta 881:268–275.
184. Reichard, P. 1988. Interactions between deoxyribonucleotide and DNA synthesis. Annu. Rev. Biochem. 57:349–374.
185. Reichard, P. 1993. From RNA to DNA, why so many ribonucleotide reductases. Science 260:1773–1777.
186. Roisin, M. P., and A. Kepes. 1978. Nucleosidediphosphate kinase of Escherichia coli, a periplasmic enzyme. Biochim. Biophys. Acta 526:418–428.
187. Roland, K. L., C. Liu, and C. L. Turnbough. 1988. Role of the ribosome in suppressing transcriptional termination at the pyrBI attenuator of Escherichia coli K-12. Proc. Natl. Acad. Sci. USA 85:7149–7153.
188. Roland, K. L., F. E. Powell, and C. L. Turnbough. 1985. Role of translation and attenuation in the control of pyrBI operon expression in Escherichia coli. J. Bacteriol. 163:991–999.
189. Roof, W. D., K. F. Foltermann, and J. R. Wild. 1982. The organization and regulation of the pyrBI operon in E. coli includes a rho-independent attenuator sequence. Mol. Gen. Genet. 187:391–400.
190. Roovers, M., D. Charlier, A. Feller, D. Gigot, F. Holemans, W. Lissens, A. Pierard, and N. Glansdorff. 1988. carP, a novel gene regulating the transcription of the carbamoylphosphate synthetase operon of Escherichia coli. J. Mol. Biol. 204:857–865.
191. Rubino, S. D., H. Nyunoya, and C. J. Lusty. 1986. Catalytic domains of carbamyl phosphate synthetase. Glutamine-hydrolyzing site of Escherichia coli carbamyl phosphate synthetase. J. Biol. Chem. 261:11320–11327.
192. Rubio, V., J. Cervera, C. J. Lusty, E. Bendala, and H. G. Britton. 1991. Domain structure of the large subunit of Escherichia coli cabamoyl phosphate synthetase. Location of the binding site for the allosteric inhibitor UMP in the COOH-terminal domain. Biochemistry 30:1068–1075.
193. Russel, M., and A. Holmgren 1988. Construction and characterization of glutaredoxin-negative mutants of Escherichia coli. Proc. Natl. Acad. Sci. USA 85:990–994.
194. Russel, M., P. Model, and A. Holmgren. 1990. Thioredoxin or glutaredoxin in Escherichia coli is essential for sulfate reduction but not for deoxyribonucleotide synthesis. J. Bacteriol. 172:1923–1929.
195. Saeki, T., M. Hori, and H. Umezawa. 1974. Kinetic studies on the inhibition of nucleoside diphosphate kinase by desdanine. J. Biochem. 76:623–629.
196. Scapin, G., C. Grubmeyer, and J. C. Sacchettini. 1994. Crystal structure of orotate phosphoribosyltransferase. Biochemistry 33:1287–1294.
197. Scapin, G., J. C. Sacchettini, A. Dessen, M. Bhatia, and C. Grubmeyer. 1993. Primary structure and crystallization of orotate phosphoribosyltransferase from Salmonella typhimurium. J. Mol. Biol. 230:1304–1308.
198. Schachman, H. K. 1988. Can a simple model account for the allosteric transition of aspartate transcarbamoylase? J. Biol. Chem. 263:18583–18586.
199. Schachman, H. K. 1993. Aspartate transcarbamoylase. Curr. Opin. Struct. Biol. 3:960–967.
200. Schachman, H. K., C. D. Pauza, M. Navre, M. J. Karels, L. Wu, and Y. R. Yang. 1984. Location of amino acid alterations in mutants of aspartate transcarbamoylase: structural aspects of interallelic complementation. Proc. Natl. Acad. Sci. USA 81:115–119.
201. Schneider, H., H. Fsihi, B. Kottwitz, B. Mygind, and E. Bremer. 1993. Identification of a segment of the Escherichia coli Tsx protein that functions as a bacteriophage receptor area. J. Bacteriol. 175:2809–2817.
202. Schwartz, M., and J. Neuhard. 1975. Control of expression of the pyr genes in Salmonella typhimurium: effects of variations in uridine and cytidine nucleotide pools. J. Bacteriol. 121:814–822.
202a. Serina, L., C. Blondin, E. Krin, O. Sismeiro, A. Danchin, H. Sakamoto, A. M. Gilles, and O. Barzu. 1995. Escherichia coli UMC kinase, a member of aspartokinase family, is a hexamer regulated by guanine nucleotides and UTP. Biochemistry 34:5066–5074.
203. Shlomai, J., and A. Kornberg. 1978. Deoxyuridine triphosphatase of Escherichia coli. Purification, properties, and use as reagent to reduce uracil incorporation into DNA. J. Biol. Chem. 253:3305–3312.
204. Smallshaw, J. E., and R. A. Kelln. 1992. Cloning, nucleotide sequence and expression of the Escherichia coli K-12 pyrH gene encoding UMP kinase. Genetics (Life Sci. Adv.) 11:59–65.
205. Søgaard-Andersen, L., and P. Valentin-Hansen. 1993. Protein-protein interaction in gene regulation: the cAMP-CRP complex sets the specificity of a second DNA-binding protein, the CytR repressor. Cell 75:557–566.
206. Sørensen, K. I. 1994. Conformational heterogeneity in the Salmonella typhimurium pyrC and pyrD leader mRNAs produced in vivo. Nucleic Acids Res. 22:625–631.
207. Sørensen, K. I., K. E. Baker, R. A. Kelln, and J. Neuhard. 1993. Nucleotide pool-sensitive selection of the transcriptional start site in vivo at the Salmonella typhimurium pyrC and pyrD promoters. J. Bacteriol. 175:4137–4144.
208. Sørensen, K. I., and J. Neuhard. 1991. Dual transcriptional initiation sites from the pyrC promoter control expression of the gene in Salmonella typhimurium. Mol. Gen. Genet. 225:249–256.
209. Spiro, S., and J. R. Guest. 1991. Adaptive responses to oxygen limitation in Escherichia coli. Trends Biochem. Sci. 16:310–314.
210. Stevens, R. C., J. E. Gouaux, and W. N. Lipscomb. 1990. Structural consequence of effector binding to the T-state of aspartate carbamoyltransferase: crystal structures of the unligated and ATP- and CTP-complexed enzymes at 2.6 Å resolution. Biochemistry 29:7691–7701.
211. Stevens, R. C., and W. N. Lipscomb. 1992. A molecular mechanism for pyrimidine and purine nucleotide control of aspartate transcarbamoylase. Proc. Natl. Acad. Sci. USA 89:5281–5285.
212. Stubbe, J. 1990. Ribonucleotide reductases. Adv. Enzymol. 63:349–419.
213. Stubbe, J. 1990. Ribonucleotide reductases: amazing and confusing. J. Biol. Chem. 265:5329–5332.
214. Sun, L., B. A. Jacobson, B. S. Dien, F. Srienc, and J. A. Fuchs. 1994. Cell cycle regulation of the Escherichia coli nrd operon: requirement for a cis-acting upstream AT-rich sequence. J. Bacteriol. 176:2415–2426.
215. Sun, X., J. Harder, M. Krook, H. Jörnvall, B.-M. Sjöberg, and P. Reichard. 1993. A possible glycine radical in anaerobic ribonucleotide reductase from Escherichia coli: nucleotide sequence of the cloned nrdD gene. Proc. Natl. Acad. Sci. USA 90:577–581.
216. Syvanen, J. M., and J. R. Roth. 1973. Structural genes for catalytic and regulatory subunits of aspartate transcarbamylase. J. Mol. Biol. 76:363–378.
217. Theisen, M., R. A. Kelln, and J. Neuhard. 1987. Cloning and characterization of the pyrF operon of Salmonella typhimurium. Eur. J. Biochem. 164:613–619.
218. Theisen, M., and J. Neuhard. 1990. Translational coupling in the pyrF operon of Salmonella typhimurium. Mol. Gen. Genet. 222:345–352.
219. Trotta, P. P., L. F. Estis, A. Meister, and R. H. Haschemeyer. 1974. Self-association and allosteric properties of glutamine-dependent carbamyl phosphate synthetase. Reversible dissociation to monomeric species. J. Biol. Chem. 249:482–491.
220. Trotta, P. P., L. M. Pinkus, R. H. Haschemeyer, and A. Meister. 1974. Reversible dissociation of the monomer of glutamine-dependent carbamyl phosphate synthetase into catalytically active heavy and light subunits. J. Biol. Chem. 249:492–499.
221. Tuggle, C. K., and J. A. Fuchs. 1986. Regulation of the operon encoding ribonucleotide reductase in Escherichia coli: evidence for both positive and negative control. EMBO J. 5:1077–1085.
222. Turnbough, C. L. 1983. Regulation of Escherichia coli aspartate transcarbamoylase synthesis by guanosine tetraphosphate and pyrimidine ribonucleoside triphosphates. J. Bacteriol. 153:998–1007.
223. Turnbough, C. L., and B. R. Bochner. 1985. Toxicity of the pyrimidine biosynthetic pathway intermediate carbamyl aspartate in Salmonella typhimurium. J. Bacteriol. 163:500–505.
224. Turnbough, C. L., K. L. Hicks, and J. P. Donahue. 1983. Attenuation control of pyrBI operon expression in Escherichia coli K-12. Proc. Natl. Acad. Sci. USA 80:368–372.
225. Turnbough, C. L., K. H. Kerr, W. R. Funderburg, J. P. Donahue, and F. E. Powell. 1987. Nucleotide sequence and characterization of the pyrF operon of Escherichia coli K12. J. Biol. Chem. 262:10239–10245.
226. Uerkvitz, W., and C. F. Beck. 1981. Periplasmic phosphatases in Salmonella typhimurium LT2. A biochemical, physiological, and partial genetic analysis of three nucleoside monophosphate dephosphorylating enzymes. J. Biol. Chem. 256:382–389.
226a. Uhlin, U., and H. Eklund. 1994. Structure of ribonucleotide reductase protein R1. Nature (London) 370:533–539.
227. Valentin-Hansen, P. 1978. Uridine-cytidine kinase from Escherichia coli. Methods Enzymol. 51:308–314.
228. Valentin-Hansen, P., B. Holst, J. Josephsen, K. Hammer, and B. Albrechtsen. 1989. CRP/cAMP- and CytR-regulated promoters in Escherichia coli K-12: the cdd promoter. Mol. Microbiol. 3:1385–1390.
229. van Alphen, W., N. van Selm, and B. Lugtenberg. 1978. Pores in the outer membrane of Escherichia coli K12. Involvement of proteins b and e in the functioning of pores for nucleotides. Mol. Gen. Genet. 159:75–83.
229a. van der Saal, W., P. M. Anderson, and J. J. Villafranca. 1985. Mechanistic investigations of Escherichia coli cytidine-5'-triphosphate synthetase. Detection of an intermediate by positional isotope exchange experiments. J. Biol. Chem. 260:14993–14997.
230. Vial, T. C., K. E. Baker, and R. A. Kelln. 1993. Dual control by purines and pyrimidines of the expression of the pyrD gene of Salmonella typhimurium. FEMS Microbiol. Lett. 111:309–314.
231. Vita, A., A. Amici, T. Cacciamani, M. Lanciotti, and G. Magni. 1985. Cytidine deaminase from Escherichia coli B. Purification and enzymatic and molecular properties. Biochemistry 24:6020–6024.
232. Vita, A., C. Y. Huang, and G. Magni. 1983. Uridine phosphorylase from Escherichia coli B: kinetic studies on the mechanism of catalysis. Arch. Biochem. Biophys. 226:687–692.
233. Vita, A., and G. Magni. 1983. One step procedure for the purification of uridine phosphorylase from Escherichia coli. Anal. Biochem. 133:153–156.
234. Vogel, U., S. Pedersen, and K. F. Jensen. 1991. An unusual correlation between ppGpp pool size and rate of ribosome synthesis during partial pyrimidine starvation of Escherichia coli. J. Bacteriol. 173:1168–1174.
235. Walter, M. R., W. J. Cook, L. B. Cole, S. A. Short, G. W. Koszalka, T. A. Krenitsky, and S. E. Ealick. 1990. Three-dimensional structure of thymidine phosphorylase from Escherichia coli at 2.8 Å resolution. J. Biol. Chem. 265:14016–14022.
236. Walton, L., C. A. Richards, and L. P. Elwell. 1989. Nucleotide sequence of the Escherichia coli uridine phosphorylase (udp) gene. Nucleic Acids Res. 17:6741.
237. Wang, L., and B. Weiss. 1992. dcd (dCTP deaminase) gene of Escherichia coli: mapping, cloning, sequencing, and identification as a locus of suppressors of lethal dut (dUTPase) mutations. J. Bacteriol. 174:5647–5653.
238. Wang, M., and G. Church. 1992. A whole genome approach to in vivo DNA-protein interactions in E. coli. Nature (London) 360:606–610.
239. Warner, H. R., B. K. Duncan, C. Garrett, and J. Neuhard. 1981. Synthesis and metabolism of uracil-containing deoxyribonucleic acid in Escherichia coli. J. Bacteriol. 145:687–695.
240. Washabaugh, M. W., and K. D. Collins. 1984. Dihydroorotase from Escherichia coli. Purification and characterization. J. Biol. Chem. 259:3293–3298.
241. Washabaugh, M. W., and K. D. Collins. 1986. Dihydroorotase from Escherichia coli. Sulfhydryl group-metal ion interaction. J. Biol. Chem. 261:5920–5929.
242. Wedler, F. C., Y. Hsuanya, E. R. Kantrowitz, and S. A. Middleton. 1989. Regulatory behaviour of Escherichia coli aspartate transcarbamylase altered by site-specific mutation of Tyr240-Phe in the catalytic chain. J. Biol. Chem. 264:17266–17274.
243. Weiss, B., and L. Wang. 1994. Synthesis de novo of thymidylate via deoxycytidine in dcd (dCTP deaminase) mutants of Escherichia coli. J. Bacteriol. 176:2194–2199.
244. Weng, M., C. A. Makaroff, and H. Zalkin. 1986. Nucleotide sequence of Escherichia coli pyrG encoding CTP synthetase. J. Biol. Chem. 261:5568–5574.
245. West, T. P., M. S. Shanley, and G. A. O’Donovan. 1982. Purification and some properties of cytosine deaminase from Salmonella typhimurium. Biochim. Biophys. Acta 719:251–258.
246. Wild, J. R., K. F. Foltermann, and G. A. O’Donovan. 1980. Regulatory divergence of aspartate transcarbamoylase within the Enterobacteriaceae. Arch. Biochem. Biophys. 201:506–517.
247. Wild, J. R., S. J. Loughrey-Chen, and T. S. Corder. 1989. In the presence of CTP, UTP becomes an allosteric inhibitor of aspartate transcarbamoylase. Proc. Natl. Acad. Sci. USA 86:46–50.
248. Wilson, H. R., C. D. Archer, J. Liu, and C. L. Turnbough. 1992. Translational control of pyrC expression mediated by nucleotide-sensitive selection of transcriptional start site in Escherichia coli. J. Bacteriol. 174:514–524.
249. Wilson, H. R., P. T. Chan, and C. L. Turnbough. 1987. Nucleotide sequence and expression of the pyrC gene of Escherichia coli. J. Bacteriol. 169:3051–3058.
250. Wilson, H. R., and C. L. Turnbough. 1990. Role of the purine repressor in the regulation of pyrimidine gene expression in Escherichia coli K12. J. Bacteriol. 172:3208–3213.
251. Xu, W., and E. R. Kantrowitz. 1989. Function of threonine-55 in the carbamoyl phosphate binding site of Escherichia coli aspartate transcarbamoylase. Biochemistry 28:9937–9943.
251a. Yamanaka, K., T. Ogura, E. V. Koonin, H. Niki, and S. Hiraga. 1994. Multicopy suppressors, mssA and mssB, of an smbA mutant of Escherichia coli. Mol. Gen. Genet. 243:9–16.
252. Yamanaka, K., T. Ogura, H. Niki, and S. Hiraga. 1992. Identification and characterization of the smbA gene, a suppressor of the mukB null mutant of Escherichia coli. J. Bacteriol. 174:7517–7526.
253. Yan, Y., and M. Demerec. 1965. Genetic analysis of pyrimidine mutants of Salmonella typhimurium. Genetics 52:643–651.
254. Yang, C., D. Carlow, R. Wolfenden, and S. Short. 1992. Cloning and nucleotide sequence of the Escherichia coli cytidine deaminase (cdd) gene. Biochemistry 31:4168–4174.
255. Zeeh, A., and D. A. Shub. 1991. The product of the split sunY gene of bacteriophage T4 is a processed protein. J. Bacteriol. 173:6980–6985.
256. Zhang, Y., and E. R. Kantrowitz. 1991. The synergistic inhibition of Escherichia coli apartate carbamoyltransferase by UTP in the presence of CTP is due to the binding of UTP to the low affinity CTP sites. J. Biol. Chem. 266:22154–22158.
257. Zhang, Y., and E. R. Kantrowitz. 1992. Probing the regulatory site of Escherichia coli aspartate transcarbamoylase by site-specific mutagenesis. Biochemistry 31:792–798.
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