Catabolism of Amino Acids and Related Compounds
LARRY REITZER
[SECTION EDITOR: AUGUST BÖCK]
Posted July 25, 2005
Department of Molecular and Cell Biology, The University of Texas at Dallas, Richardson, TX 75083-0688
Phone: (972) 883-2502 or -2524, Fax: (972) 883-2409, E-mail:
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This review considers the pathways for the degradation of amino acids and a few related compounds (agmatine, putrescine, ornithine, and γ-aminobutyrate), along with their functions and regulation. The catabolic pathways cannot be grouped into the same families as the anabolic pathways. Nonetheless, when an amino acid or related compound is degraded to another amino acid, the catabolic pathways are considered in the same section. Nitrogen limitation and an acidic environment are two physiological cues that regulate expression of several amino acid catabolic genes. These environments are considered in separate sections. This review will considerEscherichia coli, Salmonella enterica serovar Typhimurium, andKlebsiella species. The latter is included because the pathways inKlebsiella species have often been thoroughly characterized and also because of interesting differences in pathway regulation. These organisms can essentially degrade all the protein amino acids, except for the three branched-chain amino acids.
Escherichia coli, Salmonella enterica serovar Typhimurium, and Klebsiella aerogenes can assimilate nitrogen from D- and L-alanine, arginine, asparagine, aspartate, glutamate, glutamine, glycine, proline, and D- and L-serine (Table 1). There are species differences in the utilization of agmatine, citrulline, cysteine, histidine, the aromatic amino acids, and polyamines (putrescine and spermidine). Nitrogen source catabolism must generate the major intracellular nitrogen donors, glutamine and glutamate. Glutamate synthesis does not require ammonia generation for nitrogen sources that can be deaminated by transamination (e.g., aspartate) or that produce glutamate as an end product (e.g., proline or arginine), while glutamate synthesis obligatorily requires ammonia production for other nitrogen sources (e.g., serine). The former amino acids will be referred to as glutamate generating, while the latter amino acids will be referred to as ammonia generating. Regardless of the pathway of glutamate synthesis, nitrogen source catabolism must generate ammonia for glutamine synthesis. Ammonia-generating nitrogen sources must produce about four times more ammonia than the glutamate-generating nitrogen sources. Despite the importance of ammonia, the ammonia-generating reaction during the catabolism of many glutamate-generating nitrogen sources (e.g., aspartate, glutamate, proline, and putrescine) has not been identified.
Table 1Amino acids and derivatives as nitrogen sources |
The pathways of amino acid catabolism can be as quantitatively active as the pathways of carbon and energy metabolism. One gram of bacteria contains about 40 mmol of carbon and 10 mmol of nitrogen (141). Bacteria must assimilate 10 mmol of nitrogen, which is almost the same as the aerobic glucose requirement (~13 mmol), which is used for energy (~50%) and biosynthesis (~50%). To provide some indication of flux through amino acid catabolic pathways, Table 2 presents the calculated relative fluxes through the transaminases, which represent a variety of metabolic pathways. When cells are grown in glucose-γ-aminobutyrate minimal medium (i.e., γ-aminobutyrate is the sole nitrogen source), the calculated flux through the γ-aminobutyrate catabolic enzymes apparently exceeds the flux through the first half of glycolysis. Nitrogen source transport must also be quantitatively important. For example, 1 g of E. coli grown in glucose-γ-aminobutyrate minimal medium requires 10.3 mmol of γ-aminobutyrate (for nitrogen content) and an estimated 9.9 mmol of glucose. Yet transport systems for many amino acids as nitrogen sources have not been genetically identified. If transport of a nitrogen source with a single nitrogen atom requires ATP (e.g., by an ATP-binding cassette or ABC system), then synthesis of 1 g of cells would require an additional 10 mmol of ATP, which is about 14% of the total ATP requirement.
Table 2Relative velocities of transaminases |
Growth with an amino acid as nitrogen source usually results in low intracellular ammonia, which in turn results in low intracellular glutamine and induction of the nitrogen-regulated (Ntr) response (217, 410). Ntr regulators induce glutamine synthetase (which assimilates ammonia and synthesizes glutamine), a variety of transport systems for nitrogen-containing compounds (Table 3), and a few catabolic enzymes, especially those for polyamine catabolism. Ntr regulators also repress a few enzymes: AsnA, SerA, glutamate synthase, and glutamate dehydrogenase. Three functions have been proposed for the Ntr response: ammonia assimilation, nitrogen scavenging, and metabolic coordination that integrates slower growth with a variety of metabolic functions (410).
Table 3Ntr transport systems |
Strains with defects in the Ntr response, Ntr− mutants, cannot utilize most nitrogen sources, although they can still utilize putrescine and aspartate. Such Ntr mutants can result from defects in the glnA-ntrBC (or glnALG) operon, which specifies glutamine synthetase and two important regulators of the Ntr response. Failure to sense or transmit a signal of nitrogen deficiency can also prevent the utilization of a variety of nitrogen sources. This can result from null mutations in glnD (uridylyltransase/uridylyl-removing enzyme), ntrC (NtrC, also called NRI), ntrB (NtrB, also called NRII), or rpoN (σ54) and those mutations in glnB (PII) that impair uridylylation (a partial loss of function). Such mutations prevent maximal expression of the glnA-ntrBC operon from the glnAp2 promoter.
Loss of glutamate synthase (glutamine-oxoglutarate amidotransferase, or GOGAT) also prevents utilization of many organic nitrogen sources (125, 328, 368). A mutant lacking the leucine-responsive regulatory protein (Lrp), which is required for gltBD (GOGAT-encoding) expression, has a similar phenotype. Loss of integration host factor (IHF), which is also required for gltBD expression, should result in a similar phenotype, although this has not been examined (15, 377). gltBD mutants are said to have an Asm− phenotype. E. coli or Klebsiella pneumoniae gltBD mutants can still utilize asparagine, aspartate, glutamate, and glutamine because these amino acids generate glutamate. The mutants also utilize D-serine, presumably because its degradation generates ammonia so rapidly that glutamate dehydrogenase can synthesize glutamate (328, 368). Two explanations have been presented to account for the Asm− phenotype (165). Constitutive Ntr expression suppresses the Asm− phenotype. Therefore, it was proposed that without GOGAT (the major glutamine-metabolizing enzyme) glutamine accumulates and prevents Ntr gene expression (368). This can be called a glutamine-excess hypothesis. Although not usually mentioned, this suppression occurs only for growth with glutamate-generating nitrogen sources, not for ammonia-generating nitrogen sources, such as serine, alanine, glycine, asparagine, or methionine (368). The failure to suppress the defect for ammonia-generating nitrogen sources is more consistent with a glutamate-starvation hypothesis, which proposes that glutamate starvation prevents Ntr induction (165). This hypothesis assumes that there is insufficient protein turnover, and hence glutamate generation, for induction of the Ntr response. The explanation for the Asm− phenotype may depend on the type of nitrogen source, that is, the glutamine-excess hypothesis may be correct for glutamate-generating compounds and the glutamate-starvation hypothesis for ammonia-generating nitrogen sources. However, a complete explanation of the Asm− phenotype may involve other factors. For example, since glutamate and potassium pools are linked (579), glutamate starvation may deplete intracellular potassium and affect various aspects of metabolism. (A third explanation for the Asm− phenotype is that a regulatory gene cotranscribed with gltBD is required for nitrogen source utilization [64, 65]. Subsequent studies disproved this possibility [165, 169].)
Mutations that create or increase a requirement for ammonia also prevent utilization of most organic nitrogen sources. A mutation in asnB of K. aerogenes eliminates glutamine-dependent asparagine synthetase and results in cells that require AsnA-dependent asparagine synthesis, which requires ammonia. Such a mutant is unable to utilize most nitrogen sources (412). A mutation in nit/nadE of serovar Typhimurium that reduces ammonia-dependent NAD synthetase activity results in a growth requirement for ammonia and prevents utilization of a variety of organic nitrogen sources (50, 455). Finally, alterations of glutamine-dependent amidotransferases that impair amide transfer also result in a requirement for ammonia, which can replace glutamine as a substrate, and a pleiotropic nitrogen utilization defect (e.g., see reference 314).
Ornithine Decarboxylase, SpeC.
This enzyme is the primary enzyme of putrescine (1,4-diaminobutane) synthesis (Fig. 1). Purified SpeC is dimeric with a Km of 5.6 mM for ornithine (14). Putrescine and spermidine inhibit SpeC activity in vitro (14) and probably in vivo (521). Various nucleotides stimulate activity: GTP > ATP > CTP > GMP (14), whereas guanosine 5'-diphosphate-3'-diphosphate (ppGpp) inhibits SpeC (207, 208, 434). SpeC is also phosphorylated, which appears to stimulate activity (10, 280). GTP can activate unphosphorylated SpeC (10). In addition, a protein called antizyme has been proposed to bind and inhibit SpeC (280); GTP dissociates antizyme from SpeC (10). Antizyme was shown to be AtoC, a response regulator of acetoacetate catabolism (59). However, the role of this protein as an inhibitor of SpeC has been questioned (222). Several ribosomal proteins also appear to inhibit SpeC activity (241). The only reported regulation of speC expression is repression by cyclic AMP and its receptor protein (570, 571). These properties suggest an anabolic function. A catabolic function has been suggested from a single report that loss of SpeC impairs growth with ornithine as a nitrogen source (471).
Arginine Decarboxylase, SpeA.
In the presence of arginine, which represses the enzymes of ornithine synthesis, arginine decarboxylase (SpeA) and agmatine ureohydrolase (SpeB) convert arginine to putrescine (Fig. 1). (SpeB will be discussed in the section on agmatine catabolism.) Purified SpeA (Km for arginine, 30 μM) is a homotetramer (572). Putrescine and spermidine inhibit SpeA activity in vivo and in vitro (521, 572). Putrescine also represses speA synthesis (319). Cyclic AMP inhibits SpeA activity, but the effect is apparently indirect, since cyclic AMP does not affect speA transcription (from a speA-lacZ fusion) (319). Presumably cyclic AMP adjusts polyamine synthesis to the growth rate. These properties suggest an anabolic function. However, it has been reported that nitrogen limitation induces arginine decarboxylase, which suggests a catabolic function (470). Notably, a microarray analysis of Ntr genes did not suggest control by regulators of the Ntr response (587). An unusual property of SpeA is its periplasmic location, although it is not released by osmotic shock (perhaps, as suggested, because it is too large) (54). This accounts for consistent and convincing reports of separation of arginine pools from endogenous and exogenous sources and the preferential synthesis of putrescine from exogenous arginine (469, 522). In ammonia-containing minimal medium, arginine decarboxylase converts about 25% of exogenous arginine to putrescine (454, 521). However, during the slower growth with arginine as sole nitrogen source, arginine decarboxylase converts only 3% of exogenous arginine to putrescine (454). The differential conversion implies tight regulation of SpeA activity, despite its periplasmic location. Factors such as putrescine and spermidine could conceivably control the periplasmic SpeA activity. However, another possibility is that the periplasmic arginine-binding proteins, especially the nitrogen-regulated ArgT, compete with SpeA for arginine (discussed in the section on arginine catabolism).
Lysine Decarboxylase, LdcC.
Lysine decarboxylase converts lysine to cadaverine (1,5-diaminopentane), which can substitute for putrescine as a polyamine. E. coli possesses an inducible high-activity enzyme (CadA) and a constitutive low-activity enzyme. Putrescine and spermidine inhibit the basal activity in a strain in which the inducible enzyme is not expressed or in a strain with a multicopy plasmid containing the gene for the low-activity enzyme ( 558, 578). The constitutive enzyme is thermolabile in a crude extract (153). The purified low-activity enzyme is thermolabile and a decamer of 80-kDa subunits, with a pH optimum from 6 to 8 (251, 274). The gene for this enzyme was identified as ldc or ldcC (251, 578). The gene may be in an operon with a downstream gene of unknown function (578). Expression of ldc is very low and unaffected by pH or lysine (251, 274, 578). The presence of the promoter region on a plasmid moderately derepresses ldc expression, which implies a titratable but unidentified repressor (274). These results are consistent with a biosynthetic function (i.e., the synthesis of cadaverine as a polyamine).
E. colipossesses five inducible amino acid decarboxylases: AdiA (arginine), SpeF (ornithine), CadA (lysine), GadA (glutamate), and GadB (glutamate). They participate in acid survival, and some can amount to 7% of the total cellular protein (520). The phenotypes of mutants lacking lysine or glutamate decarboxylase grown in neutral pH medium (discussed subsequently) may indicate additional functions.
Arginine Decarboxylase, AdiA.
Gale first observed induction of an arginine decarboxylase in E. coli (144). The enzyme is induced in anaerobic, acidic media with a casein hydrolysate (arginine, methionine, tyrosine, and asparagine mimic the casein hydrolysate) (Fig. 1) (306). Induction also requires iron (306) for reasons that have not been investigated. When induced, the enzyme accounts for ~0.3% of total cellular protein (41). The purified protein from E. coli B attacks arginine (Km, 0.65 mM), has an acidic pH optimum, contains pyridoxal 5'-phosphate, and has 10 identical subunits (41, 42). The purified inducible decarboxylase is about 30 times more active than purified SpeA (572). The adiA gene codes for the inducible arginine decarboxylase (21, 507), and gene expression parallels protein synthesis (21). Expression is enhanced by IHF, CysB, and AdiY (a XylS/AraC-like regulator) and is repressed by H-NS (reviewed in reference 426). AdiA-mediated arginine degradation permits survival of E. coli at pH ≤ 2.5 (63). Adi-dependent arginine decarboxylation absorbs an intracellular proton and removes an acidic group, which is released as CO2. (Notice that the periplasmic SpeA would consume an extracellular proton.) The AdiC protein is an arginine-agmatine antiporter that removes the product of decarboxylation (162, 224). The three adi genes are clustered: adiAY forms an operon, while the downstream adiC is monocistronic (162).
Ornithine Decarboxylase, SpeF.
The inducible ornithine decarboxylase (SpeF) is optimally expressed anaerobically in acidic medium (pH ≤ 5.5) with at least 0.8% ornithine (Fig. 1) (13, 244, 487). A 14-fold purification to homogeneity from a crude extract suggests that SpeF might account for up to 7% of the total cellular protein (13). The Km for ornithine is 3.6 mM, and putrescine or spermidine moderately inhibits it (13). Like the biosynthetic ornithine decarboxylase, nucleotides stimulate activity of the inducible enzyme: GTP > UTP > ATP > CTP (14). The proximity of speF to potE (which codes for an ornithine-putrescine antiporter) suggests that they form an operon (244). In other words, the decarboxylase and antiporter are functionally linked, as is the case for AdiA and AdiC. The factors that control speF-potE expression are not well characterized, and an ornithine-responsive regulator has not been identified. Loss of the activator of the cadBA system (see following) increases speF-potE expression, suggesting reciprocal regulation (487). RNase III removes part of the 5'-untranslated region, which increases translational efficiency (246).
Lysine Decarboxylase, CadA.
The inducible lysine decarboxylase (CadA) is optimally expressed anaerobically in complex acidic medium with at least 0.8% lysine (432). A mixture of nine amino acids could replace the nutrient broth, and some amino acids (e.g., alanine) were inhibitory (61). Specific activity decreases in stationary phase (389, 432). The purified enzyme is a decamer and is highly specific for lysine (Km, 1.5 mM) (432). The cadA gene is part of a cadBA operon, which has one promoter preceding cadB (307, 308, 556). CadB is a cadaverine transporter at neutral pH, and a lysine-cadaverine antiporter at acidic pH (487). Not surprisingly, it is induced by the same factors that induce CadA (487).
A variety of factors has been implicated in cadBA expression (Fig. 2). CadC is an activator of cadBA expression (334, 556). The cadC gene is just upstream from the cadBA operon (556), and cadC expression appears to be constitutive (335). CadC mediates both pH- and lysine-dependent induction (107, 334). Cadaverine represses cadBA, and CadC mediates this repression (334). Perhaps the ratio of lysine to cadaverine controls CadC activity. CadC is localized in the membrane, which is unusual for a transcriptional activator (107). Alterations that result in either pH or lysine independence are generally in the periplasmic domain (107). CadC appears to sense pH and lysine separately (335). The cadR/lysP locus also contributes to induction by lysine (334, 389). Mutations in cadC are epistatic to those in lysP (334). LysP is one system that transports lysine (500); the LAO system is the second lysine transport system (described in the section on arginine catabolism). An interaction between LysP and CadC has been proposed to mediate the induction by lysine (107, 334, 335). Lysine represses lysP expression, which may contribute to induction by lysine; however, the activity of LysP is probably also important for cadBA expression (335). In addition to these factors, H-NS represses cadBA during aerobic growth (475), but an hns mutant still exhibits pH-dependent induction of cadBA (409); and the ArcA and ArcB two-component system appears to be required for cadBA expression (409). Control by hns appears to be epistatic to the arc regulators (409). Sodium carbonate inhibits cadBA expression, and lysine decarboxylase appears to generate CO2 when oxygen is limiting (524). The control by CO2 was proposed to account for the transient cadBA expression during inducing conditions (335).
An unanticipated function of cadA is regulation of cell division (400). CadA appears to inhibit cell division (a cadA mutant continues to divide at a time when wild-type cells enter stationary phase) (400). FlhD, a regulator of flagellar gene expression, activates cadBA expression independently of CadC (400). This activation apparently requires depletion of serine from a complex medium and acetyl phosphate-dependent phosphorylation of OmpR (400).
The Glutamate Decarboxylases, GadA and GadB.
E. colihas two glutamate decarboxylases, GadA and GadB (101, 482). The purified proteins are homohexamers (60, 508, 529, 530). The two isozymes differ by only five amino acid residues and have virtually identical properties (101). It seems likely that earlier preparations contained both enzymes and that the reported properties are from a mixture of the two proteins. The purified enzyme has a Km for glutamate of 0.8 mM, but the apparent Km determined from crude extracts can be as high as 10 mM (286, 329, 481). Chloride ions stimulate decarboxylase activity, while some citric acid cycle intermediates (citrate, cis-aconitate, and α-ketoglutarate) competitively inhibit decarboxylase activity (148, 481). Glutamate decarboxylase has an acidic pH optimum (481), which is consistent with a function during survival in extreme acid environments (~pH 2). In the transition from a neutral to an acidic environment (pH ≤ 5), a conformational change alters the absorption spectrum of the enzyme and activates decarboxylase activity (60; reference 360 and references therein). Chloride stabilizes the active form of the enzyme (360). At neutral pH, the enzyme is cytoplasmic (92%), but at acidic pH, it is sequestered to the membrane (~55%) (60). Presumably, the association with the membrane further stabilizes the active form of the protein.
The glutamate decarboxylases in E. coli, the products of gadA and gadB, have been intensively studied because their presence helps cells survive in extremely acidic environments (pH 2 to 3 for several hours) (e.g., during passage through the stomach). (A complete discussion of acid resistance will be presented in a future EcoSal chapter.) However, the slower growth of a gadA gadB double mutant in minimal medium at neutral pH (236) and induction of GadA and GadB in high pH media (39, 43, 497, 583) suggest further functions. When induced, GadA and GadB appear to contribute equally to decarboxylase activity (100). Loss of both, but not either enzyme singly, reduces resistance to extreme acid (63) and moderately impairs growth in minimal medium (236). The response to acidity involves import of extracellular glutamate, proton-consuming glutamate decarboxylation, and export of the resulting GABA. Glutamate-GABA exchange catalyzed by GadC is absolutely required for glutamate-dependent acid resistance (63, 100, 198, 302) and is obligatorily coupled to chloride export (223).
It has been suggested that a futile cycle results if GadC transports protonated glutamate into the cell, which releases a proton upon entry that is then consumed during decarboxylation (63). This cycle ignores GABA, which is protonated when it exits the cell. It would be more accurate to state that transport of protonated glutamate and subsequent decarboxylation cannot help neutralize cytoplasmic acidity, whereas transport of unprotonated glutamate with the subsequent proton-consuming decarboxylation reaction could.
The gadA gene is part of a gadAX operon, and gadB is part of a gadBC operon. Each operon contains a single promoter preceding the first gene, and there appears to be a promoter preceding gadX (62, 100, 533, 536). Figure 3 summarizes regulatory proteins and some environmental factors that control gadAX and gadBC expression. The regulation of these two operons is complex, not only because of the number of regulatory factors, but also because of contradictory results. There is general agreement that GadE is an essential regulator and that other regulators affect either its level or activity (209, 287, 535). There is no agreement, however, on which regulators control GadE activity. It has been reported that YdeO, but not GadX, activates GadE synthesis (302). The converse (YdeO-independent, GadX-dependent expression) has also been reported (287, 536). GadX and GadW modify GadE activity, and overexpressed GadE overcomes this regulation, which suggests that GadE is limiting (287, 288, 536). GadX and GadW are DNA-binding proteins that may facilitate binding of GadE to DNA (287, 288, 476, 533).
Another regulatory circuit requires the DNA-binding response regulator EvgA, which acts through YdeO ( 301, 302). GadE is a LuxR homolog, while GadX, GadW, and YdeO are AraC homologs. There appear to be no precedents for AraC-like proteins stimulating the activity of a LuxR-like protein. A variety of other regulatory and environmental factors affect these central regulators, and they may act at multiple sites. GadA or GadB synthesis (or gadA or gadB expression) requires σS in cells grown to stationary phase at neutral pH in minimal or rich media (62, 63, 287, 288). H-NS represses one or more component of the regulatory cascade (100, 210, 288, 301, 533), although there is a report to the contrary (236). Cyclic AMP receptor protein (CRP) also represses components of the regulatory cascade (62, 236, 288). Finally, there have been isolated reports of additional regulatory or environmental factors that influence GadAB synthesis. For example, acetate induces expression (16), polyamines may be required for expression (236), and trimethylamine N-oxide complexed to TorR represses in an anaerobic environment (43). Loss of MppA, a periplasmic protein that binds a murein tripeptide results in massive overproduction of GadA and GadB (276), and SlyA overproduction induces GadA (494).
Genomic and Proteomic Analyses.
The results from numerous analyses of microarray or two-dimensional polyacrylamide gels show that acidic environments or overexpression of regulators that induce several decarboxylases has no effect on other amino acid catabolic pathways (209, 301, 302, 494, 535, 536). Therefore, the products of decarboxylase reactions should be exported without competition from other pathways. However, there is at least one situation in which the product of a decarboxylase, γ-aminobutyrate (GABA), is present with a relevant catabolic pathway, the GABA pathway (which is described below). GadA protein is synthesized at a higher level in an anaerobic alkaline environment than in an acidic environment (39, 583). High pH also appears to induce GabT (a catabolic GABA transaminase) and AstD (a dehydrogenase in the major arginine catabolic pathway) (497). It was suggested that GABA could ameliorate cytoplasmic alkalinization by generating succinate or other membrane nonpermeant acids (583). However, the properties of purified glutamate decarboxylase appear to suggest that GadA is inactive at high pH. In contrast, the AdiA and AST pathways appear to be reciprocally regulated. AdiA is induced in an anaerobic acidic environment (described above), whereas the AST pathway is induced in an alkaline environment (497).
E. coli, serovar Typhimurium, and K. aerogenes can utilize L- and D-alanine as carbon or nitrogen sources. The catabolic pathway is the same for either carbon or nitrogen source utilization. Catabolism of exogenous L-alanine involves transport, racemization to D-alanine, and oxidative deamination by D-amino acid dehydrogenase to pyruvate and ammonia (Fig. 4). L-Alanine is the only L-amino acid catabolized through its D-isomer (304). The genes and enzymes of L- and D-alanine catabolism are intricately associated with those of D-alanine synthesis and are more thoroughly described in Chapter Biosynthesis of Glutamate, Aspartate, Asparagine, l-Alanine, and d-Alanine, which should be consulted for reference. An unexpected aspect of this pathway is that the alanine transaminases (i.e., glutamic-pyruvic transaminases) do not participate in alanine catabolism. This suggests that these transaminases do not catalyze reversible reactions in vivo.
The CycA protein transports D-alanine, D-serine, glycine, and cycloserine; it is a secondary transport system for L-alanine. CycA also has recently been shown to be the major transport system for β-alanine (458). Transport is coupled to the proton motive force. A cycA mutant is unable to utilize D-alanine as a carbon source. Utilization of D-alanine as a nitrogen source was not examined but presumably also requires cycA. Expression of cycA is induced by L- or D-alanine, carbon limitation, or nitrogen limitation. Nitrogen limitation causes a three- to eightfold induction of expression and requires a transcriptional activator that is induced by nitrogen limitation, the nitrogen assimilation control protein (Nac) (587). Ntr control is alanine independent, because alanine is not required for induction (587). Alanine as a nitrogen source only minimally activates nac expression, which further indicates that control is alanine independent (B. Schneider and L. Reitzer, unpublished observation). In contrast, nitrogen sources, such as arginine, induce Nac synthesis effectively. These control mechanisms are not obviously consistent with a catabolic function for CycA during nitrogen limitation but perhaps are consistent with a modulatory function, that is, to lower intracellular alanine pools during slow growth.
The LIV-I transport system (livJ and livHMGF operons) for branched-chain amino acids transports L-alanine. Lrp without leucine (or alanine) activates liv operon expression, while Lrp with leucine (or alanine) represses expression. This regulation is most consistent with a scavenging function. There may be other physiologically relevant L-alanine transport systems, since neither transport mutants that cannot utilize L-alanine (as carbon or nitrogen source), nor L-alanine utilization in LIV-I-deficient mutants has been examined.
E. coliand serovar Typhimurium contain two alanine racemases. The racemases from serovar Typhimurium have been studied more extensively. Alr is a constitutive, low-activity alanine racemase. DadX in E. coli (called DadB in serovar Typhimurium) is an alanine-inducible high-activity racemase. Both serovar Typhimurium enzymes contain pyridoxal phosphate and were initially proposed to be monomeric (126, 554). However, recent genetic evidence indicates that the racemases are functionally dimeric (509), and biochemical evidence suggests that protein concentration and substrates affect dimerization (584). The dadX/dadB gene is part of a dadAX (dadAB) operon and dadA specifies a subunit of D-amino acid dehydrogenase. Mutations in this operon prevent L- or D-alanine catabolism and, thus, DadX/DadB is the major anabolic and catabolic racemase.
D-Amino acid dehydrogenases purified from E. coli B and assayed from crude extracts of E. coli K-12 have similar, but not identical, substrate specificities. Both enzymes deaminate D-alanine, D-phenylalanine, D-methionine, and D-α-aminobutyrate (134, 361). The dehydrogenase is membrane associated and donates electrons to the electron transport chain (181, 361, 362). The E. coli B enzyme is an iron-sulfur flavoprotein comprising two subunits with molecular weights of approximately 45,000 and 55,000 (361). The dadA gene specifies the small subunit and is in an operon that also specifies the inducible racemase. Seemingly convincing genetic evidence indicated that the structural gene for the large subunit might be localized between the ara and leu operons (29, 134, 135). However, deletion of all the genes between these operons does not affect growth with either alanine isomer (N. Gurich-Vrasic and L. Reitzer, unpublished observation). Therefore, the gene for the large subunit has yet to be identified.
The control of dad operon expression is described more fully in Chapter Biosynthesis of Glutamate, Aspartate, Asparagine, l-Alanine, and d-Alanine. In brief, cyclic AMP and CRP activate expression of the dad operon, and loss of CRP prevents utilization of alanine as a carbon source. Alanine induces dad operon expression and Lrp with alanine (or leucine) activates expression of the dad operon, while Lrp without alanine (or leucine) represses expression. Finally, Nac activates dad expression in K. aerogenes, but not in E. coli or serovar Typhimurium (225). A strain with a defect in dadQ (also called alnR) fails both to synthesize the dehydrogenase and to utilize alanine as a carbon source (29, 135). The dadQ gene maps to about minute 99 of the E. coli chromosome, but it has not been definitively identified, and its function is unknown.
The pattern of regulation suggests that the enzymes of L- or D-alanine catabolism are induced by slow growth resulting from carbon limitation, nitrogen limitation, or guanosine tetraphosphate (which controls Lrp synthesis). If so, then an important function of these enzymes (besides the obvious function of alanine catabolism) may be to lower the ratio of L-alanine to pyruvate, which might stimulate pyruvate-requiring reactions. Such a modulatory function is also consistent with the complex effects of Lrp. For example, Lrp with alanine (or leucine) represses the transporter CycA but activates the dad operon. L-Alanine induces the Ntr response even in ammonia-containing minimal medium, probably because of its inhibition of glutamine synthetase (L. Reitzer, unpublished observation). Ntr mutants fail to utilize alanine as a nitrogen source, even though Ntr regulators have little effect on expression of alanine catabolic genes. Therefore, alanine catabolism may prevent alanine toxicity.
Substantial progress has been made in understanding the catabolism of these metabolically related compounds in E. coli since the most recent reviews (304, 411). Figure 5 shows the metabolic relationships between these compounds. E. coli, serovar Typhimurium, and K. aerogenes exhibit strikingly different utilization patterns. E. coli can utilize each as sole nitrogen source, but none as sole carbon source. K. aerogenes can degrade each as sole nitrogen or carbon source. Serovar Typhimurium can catabolize arginine but none of the other four compounds as a nitrogen source and none as sole carbon source. Only some of these differences are understood.
Bacteria contain a variety of pathways for arginine catabolism, which can be used to supply nitrogen, energy, or carbon (95). The products of arginine transport and a preliminary genetic analysis suggested an arginine decarboxylase pathway for E. coli arginine catabolism (471, 563). However, an important prediction of this pathway, stoichiometric urea generation, was not met for either E. coli or K. aerogenes (140, 454). Instead, the only arginine catabolic pathway that is active during growth of E. coli, serovar Typhimurium, and probably of K. aerogenes on arginine is the arginine succinyltransferase (AST) pathway (Fig. 5), which degrades arginine as a nitrogen source (all three organisms) or as a carbon source (K. aerogenes only).
The intermediates of the AST pathway, succinylarginine, succinylornithine, succinylglutamic semialdehyde, and succinylglutamate, are similar to those of arginine synthesis, except that the former are succinylated, whereas many of the latter are acetylated. AstC is the only enzyme of the pathway that has been characterized beyond assays from crude extracts (454, 455, 496). AstC is the previously described arginine-inducible ArgM, which at high levels can suppress a defect in the biosynthetic argD (reference 418 and references therein). AstC/ArgM has been purified from E. coli (130, 454). The catabolic N2-acetylornithine 5-aminotransferase purified from K. aerogenes is undoubtedly AstC (138). In both organisms, this enzyme has the highest affinity for succinylornithine as a substrate (95, 496), but it can also deaminate acetylornithine and ornithine (38, 138). Although other AST enzymes have not been biochemically characterized, the structures of AstA and AstB have been predicted (477) and the structure of AstB determined (531a).
The astCADBE operon in E. coli specifies all five AST pathway enzymes in E. coli (454). Loss of this operon prevents growth with arginine as the nitrogen source, impairs growth with ornithine, and impairs survival during carbon starvation in E. coliand serovar Typhimurium (131, 284, 454). Nitrogen limitation induces AST pathway enzymes and elevates the transcripts for these genes (284, 454, 587). The E. coli astCADBE operon contains two promoters. Transcription from an Ntr-dependent promoter requires σ54 and phosphorylated NtrC (258, 284). In E. coli, arginine induces operon expression about threefold better than glutamine and this induction requires ArgR, the arginine repressor (258). This induction has been called superinduction, since ArgR is not required for expression in vivo or in vitro (258). In contrast, expression of the serovar Typhimurium ast operon absolutely requires ArgR in vivo, which implies that arginine is required for induction, since ArgR binds DNA only in the presence of arginine (284). In both organisms, ArgR complexed with arginine binds between the sites for NtrC and RNA polymerase and presumably promotes a protein-protein interaction (258, 284). Carbon starvation (in medium with low glucose) induces the serovar Typhimurium ast operon, and this induction required cyclic AMP, CRP, and ArgR (284). Carbon limitation (in glycerol-ammonia minimal medium) does not induce the E. coli ast operon (258). However, the E. coli ast operon was transcribed in stationary phase cells grown in broth or some glycerol-containing minimal media (258). This transcription initiates from a σS-dependent promoter that is 5 bases downstream from the Ntr promoter (258). Expression from the two E. coli promoters is competitive (131, 258). Purified σS-containing RNA polymerase was sufficient for transcription from the σS-dependent promoter, and CRP-cyclic AMP, or ArgR-arginine, or both did not affect this transcription (258). Furthermore, CRP (with or without cyclic AMP) did not bind specifically to the ast promoter region (258). The cell-free culture medium left after growth in broth also induces the ast operon (23), and indole was shown to be the active factor (552). Finally, an alkaline environment appears to induce AstD and probably other AST enzymes (497). These latter two responses might be related, since indole is the product of tryptophanase, which is also induced in alkaline media (583). The mechanism of indole- or alkaline-activated gene expression is not known. The function of such expression in cytoplasmic acidification is discussed at the end of the amino acid decarboxylase section.
In an early study of arginine transport, ion exchange chromatography separated three periplasmic binding proteins (PBPs) for arginine: a lysine-arginine-ornithine (LAO) protein, ArgBP-I, and ArgBP-II (Table 4) (73, 74, 422). HisJ, the histidine PBP, also binds arginine (Kd, 10 μM) (6). The LAO protein is the product of argT, which is within the argT-hisJQMP region (268). The LAO/ArgT protein binds lysine, arginine, and ornithine with dissociation constants of 3.0, 1.5, and 5.0 μM, respectively (422). Loss of ArgT in serovar Typhimurium reduces arginine binding by periplasmic proteins from nitrogen-limited cells by about 50% but has no effect on growth with arginine as a nitrogen source (268). Similarly, a hisJ mutant grows normally with arginine (266). A hisJ argT double mutant grew very poorly on arginine as a nitrogen source, which suggested that both proteins transport arginine during nitrogen-limited growth (268). HisP is the nucleotide-binding membrane component for HisJ and ArgT (202, 317), and its loss eliminates growth with arginine as a nitrogen source in a wild-type strain and reduces growth in a strain with a constitutive Ntr system (266, 268). Perhaps an artificially high level of another membrane component of an ABC system under Ntr control can substitute for HisP. A hisP mutant retains high-affinity arginine transport (266), which presumably involves either ArgBP-I or ArgBP-II. Nitrogen limitation induces the LAO system in E. coli and serovar Typhimurium, and this induction does not require arginine (268, 587). Transcription appears to initiate from a single σ54-dependent promoter preceding argT (411, 453). A single NtrC-binding site has been identified, but it is not clear whether it is involved in transcriptional control (8, 453). The LAO protein is also present in cells that are not nitrogen limited (75, 424). Furthermore, growth into stationary phase and carbon limitation induce argT expression (8, 453, 502). The proteins and DNA sites that mediate this control are unknown. The argT-hisJQMP region and its regulation are more thoroughly discussed in the section on histidine catabolism.
Table 4Arginine and putrescine transport systems |
ArgBP-I is the product of artJ (564). ArtJ/ArgBP-I specifically binds arginine (Kd, ~0.4 μM), but not ornithine (564). Overproduction of ArtJ increases arginine transport twofold (564).
Originally, the structural gene for ArgBP-II, which was called abpS, was thought to map near argA (67, 68), which is at minute 63.5 of the current E. coli map. In lieu of a more precise location, the published amino acid composition of ArgBP-II (70, 423) was used to identify the gene (411), which unexpectedly demonstrated that artI encoded ArgBP-II. The artI gene is located at minute 19.4 (411). This assignment is consistent with the observation that ArtI purifies with ArgBP-II (564). The properties of this protein are unusual. ArgBP-II binds arginine (Kd, 0.2 μM), but not ornithine, at 4° (67, 422). It binds both arginine (Kd, 0.2 μM) and ornithine (Kd, 10 μM) at 22° and was therefore called the AO protein (67). The elevated arginine and ornithine transport in mutants with elevated ArgBP-II/AO is consistent with the binding of both substrates in vivo (73).
The two arginine PBPs, ArtJ and ArtI, are specified by genes in the artPIQM-artJ locus. Although adjacent, artPIQM and artJ form separate transcription units and appear to be differentially regulated (564). ArgR sites have been identified by computer analysis near the artP and artJ promoters (318). However, a missense mutation in argR had no effect on arginine transport, and arginine in minimal medium had no effect on ArtI synthesis (73, 564). These results argue against arginine-specific control. However, a few observations have been presented about ArtI synthesis; growth in nutrient broth represses, acetate moderately induces, and formate moderately represses (257, 564). In contrast, arginine substantially reduced artJ expression, and lysine substantially increased artJ expression (564). Conversely, growth of E. coli (a major cause of urinary tract infections) with urine is arginine limited, and such growth induces artJ transcription about 17-fold (431). The response of artJ expression to changes in environmental arginine suggests that its function is anabolic.
A curious feature of the arginine PBPs is that ArgBP-II/AO and LAO are phosphorylated in vivo and in vitro (69, 70). Osmotic shock releases the kinase, which implies that it is either periplasmic or, more likely, loosely associated with the membrane (70). The kinase is the product of argK (72). Lesions in argK result in the cells having defective arginine transport, even though the amount of PBPs and their binding affinities for arginine are unaffected (69). However, the kinase activity of ArgK is not required for transport (72). Instead, ArgK has a substrate-independent ATPase activity (541), which appears to be required for transport (72). It has been proposed that ArgK couples ATP hydrolysis to transport (72), which echoes a suggestion made 30 years ago (424). However, this seems to conflict with the properties of ABC systems, and specifically with the ArgBP-II/AO and LAO systems, which already contains an energy-transducing ATPase, that is, HisP for LAO-dependent transport (at least when arginine is the nitrogen source), and presumably ArtP for ArtI and ArtJ. Direct evidence for an energy-transducing function for ArgK is currently not available.
Resistance to canavanine, a plant-derived antimetabolite, has been a frequent tool in the analysis of arginine transport. One gene for canavanine resistance was mapped to a locus called argP at minute 66 of the current E. coli chromosome (74, 289, 424). These mutants exhibit defective transport by the ArgBP-II/AO and LAO systems, which suggests that they transport canavanine (73, 74, 424). However, the defect in these mutants is not obvious, since their PBPs are quantitatively and qualitatively normal. A different canavanine-resistant mutant has a defect in argK (b2918), which is tightly linked to, but distinct from, argP (b2916) (69, 72). It has been proposed that ArgP controls argK expression (71; but see reference 333).
A recent genetic analysis of argP has clarified its role in canavanine resistance (333). An argP null mutant is not resistant to canavanine but is instead hypersensitive (333). Insertions in yggA (renamed argO, also called b2923), which is linked to argP, also result in hypersensitivity to canavanine (333). ArgO appears to be an arginine exporter, which excretes arginine at a high level and confers resistance to canavanine (333). Arginine secretion in a canavanine-resistant mutant had been noted 30 years ago (424). In addition to canavanine secretion, it has been suggested that ArgO might prevent toxic arginine accumulation, maintain a balance of arginine and lysine, or both. Since ArgP is a LysR-type regulator that activates argO expression (333), it was proposed that the previously isolated canavanine-resistant mutants resulted in a constitutively active ArgP and argO overexpression, which explains the trans-dominance of argP in these mutants (71, 333). ArgP-dependent expression requires intracellular arginine and is antagonized by intracellular lysine (71, 333). The antagonistic effects of arginine and lysine are also seen in control of ArtJ synthesis, which might suggest ArgP-dependent control. Most remarkably, ArgP is the previously described IciA protein that binds the origin of replication, inhibits DNA replication initiation, and also activates dnaA and nrd expression (187, 212, 213, 272, 528). It is not known whether arginine or lysine affects these activities. IciA/ArgP synthesis is induced by entry into stationary phase and phosphate limitation, but not by carbon starvation, the stringent response, or the SOS response (188, 214). Overproduction of this protein results in a growth lag upon transfer to fresh medium (528). However, ArgP is clearly not an essential protein (333, 528).
K. aerogenes, unlike E. coli, can use arginine as a carbon source. To gain insight into the requirements for utilization of an amino acid as a carbon source, astCADBE expression in K. aerogenes and E. coli was compared (454). Nitrogen limitation, at least with glutamine as the nitrogen source, induces the ast operon in these two organisms to the same levels, and this induction requires NtrC. However, if arginine is in the medium, the ast operon in K. aerogenes is induced to levels that are higher than could be achieved by E. coli. This occurs regardless of nitrogen availability or the mutational loss of NtrC. With arginine as the carbon source, the level of ast operon expression is 25 times higher than the fully induced level during nitrogen-limited growth. This makes sense, because synthesis of 1 g of E. colirequires 80 mmol of carbon (half for energy and half for biosynthesis) or about 13 mmol of arginine if it could be used as the sole carbon source. However, the 10-mmol nitrogen requirement can be fulfilled with 2.5 mmol of arginine. Therefore, utilization of arginine as a carbon source appears to require a strong and properly regulated promoter, which E. coli does not possess. Undoubtedly, use of arginine as a carbon source also requires a more active transport system than is provided by those described above.
Although pathways of putrescine catabolism in microorganisms have been reviewed (270), enormous gaps exist in our understanding of putrescine catabolism in bacteria. E. coli mutants unable to utilize putrescine as a nitrogen source have not been isolated. Furthermore, serovar Typhimurium has all the known putrescine catabolic genes but cannot utilize putrescine as a nitrogen source. Much of our current understanding of the proposed pathway, genes, and enzymes of putrescine catabolism is not based on genetic analysis but instead on assays of enzymes from extracts of wild-type strains and mutants with enhanced catabolic activity (252, 392).
Some strains of E. coli can utilize agmatine as a nitrogen source (456, 470, 471), whereas others can not (438). Agmatine ureohydrolase cleaves agmatine to putrescine and urea. K. aerogenes can subsequently degrade urea, but E. coli and serovar Typhimurium can not. The speB gene encodes the only known agmatine ureohydrolase in E. coli (518). A speB missense mutant grows less well with agmatine as a nitrogen source, but it still grows (471). A deletion of the speB gene has not been introduced into a strain that can utilize agmatine.
The E. coli agmatine ureohydrolase has been purified (agmatine Km, ~1.5 mM) (439). The enzyme is inhibited by ornithine (Ki, ~6 mM) and arginine (Ki, ~9 mM), but neither is present in cells at these concentrations (439). Polyamines do not affect activity (439). Arginine and agmatine activate speB expression threefold; arginine generates agmatine, which is probably the actual inducer (438, 470). Cyclic AMP represses expression twofold, which putrescine reverses (438); the cyclic AMP control appears to be indirect (519). It has also been reported that cyclic AMP activates expression (470). The differences in regulation may reflect strain differences. Considering that putrescine content is inversely correlated with growth rate (537), cyclic AMP-mediated repression seems more likely. In addition to these controls, the product of yggG, which is just downstream from speB and convergently transcribed, has been proposed to be a posttranscriptional regulator of speB (519). speB is the second gene of the complex speAB operon. speB can be transcribed either from a promoter preceding speA or from a possible promoter preceding speB (518, 519). Agmatine activates expression from the speB promoter (519). It has been reported that nitrogen limitation induces agmatine ureohydrolase activity in E. coli and K. aerogenes (139, 470); however, a microarray analysis of Ntr genes in E. coli did not directly implicate Ntr regulators in such control (587).
Putrescine can be degraded to succinate in four steps. A transaminase or an oxidase can catalyze putrescine deamination to γ-aminobutyraldehyde, but only the former has been identified in E. coli (253, 270, 392). ygjG specifies this enzyme (402, 437), and it is probably the previously described pat gene (471). A ΔygjG/pat mutant has lost all putrescine transaminase activity but unexpectedly grows slightly faster with putrescine as a nitrogen source (C. Pybus, B. L. Schneider, and L. Reitzer, unpublished observation). Purified putrescine transaminase specifically deaminates putrescine (Km, 9.2 mM) and cadaverine but not other substrates of so-called ω-transaminases (402, 437). K. aerogenes appears to contain a putrescine-specific transaminase and a broader specificity transaminase that can deaminate either putrescine or GABA (139). Nitrogen limitation induces putrescine transaminase activity and elevates ygjG/pat transcripts (139, 402, 470, 587).
An NAD-dependent dehydrogenase oxidizes γ-aminobutyraldehyde to GABA (252, 391, 392, 471). A mutant with eightfold less dehydrogenase activity grew normally with GABA, less well with arginine, ornithine, and putrescine, but not at all with Δ-1-pyrroline (a cyclized form of γ-aminobutyraldehyde) (471). The basis for defective arginine utilization is not obvious. The mutation in this strain was mapped between minutes 28 and 32 (471), but the gene specifying γ-aminobutyraldehyde dehydrogenase has not been identified. It has been reported that nitrogen limitation induces dehydrogenase activity (470). A γ-aminobutyraldehyde dehydrogenase has been purified and characterized from an E. coli mutant that can utilize putrescine as a carbon source (Kms of 31 and 54 μM for γ-aminobutyraldehyde and NAD, respectively). The enzyme is a dimer with a subunit Mr of 95,000 ± 1,000 (391). Whether this enzyme participates in γ-aminobutyraldehyde oxidation in a wild-type strain has not been established.
The last two steps of putrescine degradation are the same as those in GABA degradation: deamination of GABA and oxidation of succinic semialdehyde. Two different transaminases deaminate GABA: GabT and an unidentified putrescine-inducible enzyme (456). The latter accounts for about 25% of total GABA transaminase activity (456). At least two dehydrogenases oxidize succinic semialdehyde. One is GabD and the other has yet to be identified, but it is inducible by putrescine (456). The latter accounts for about 40% of the total dehydrogenase activity (456). Loss of GabT or GabD (or both) has no effect on growth rate or growth yield with putrescine as the nitrogen source, which underscores the importance of the unidentified putrescine-inducible enzymes (456). The two putrescine-inducible enzymes are not under Ntr control (456).
E. colicontains a surprising number of polyamine and putrescine transport systems (Table 4). Kazuei Igarashi and colleagues have studied three putrescine transport systems intensively (reviewed in references 215 and 216). The potABCD system specifies an ABC (ATP-binding cassette) transporter that transports spermidine better than putrescine. PotD is the PBP component that binds spermidine (Kd, 3.2 μM) with a higher affinity than putrescine (Kd, 100 μM) (242). Convincing evidence has been presented that a cytoplasmic form of PotD is also a transcriptional repressor (12). The potFGHI system specifies an ABC transporter that specifically transports putrescine. The PBP component is PotF (putrescine Kd, 2.0 μM) (381). The potE system consists of a putrescine/ornithine antiporter that can transport putrescine bidirectionally (243). Putrescine excretion has been proposed to be an important factor in maintaining putrescine homeostasis (240), and it was proposed that PotE excretes putrescine (243). However, putrescine is still excreted in a potE mutant after osmotic shock (448). The potE gene is part of the speF-potE operon, is expressed in an acidic environment, and requires ornithine for induction (13, 244). In addition to these systems, the products of ydcSTUV are orthologs to the proteins of the potABCD and potFGHI systems. Deletion of any of these four systems has no effect on growth with putrescine as a nitrogen source (B. L. Schneider and L. Reitzer, unpublished observation). Nitrogen limitation induces potFGHI and ydcSTUV (587).
A relationship between nitrogen limitation and putrescine metabolism is clearly established. Nitrogen limitation induces two putrescine transport systems, YgjG, γ-aminobutyraldehyde dehydrogenase activity (although the gene for this activity has not been identified), GabT, and GabD. However, loss of the regulators of the Ntr response has little effect on utilization of putrescine as a nitrogen source (456) (A. Kiupakis, B. L. Schneider, and L. Reitzer, unpublished observation). These results raise the issue of the role of the Ntr system in putrescine catabolism. Perhaps the Ntr regulators lower the polyamine content during the slower nitrogen-limited growth and thus modulate the activities of processes affected by polyamines (456).
E. coliK-12 can use γ-aminobutyrate (GABA) as a nitrogen source (456), despite a previous report to the contrary (117). Mutants can be isolated that utilize GABA as sole carbon source (117). Loss of the repressor GabC, also called CsiR, permits such growth (456). Wild-type K. aerogenes can use GABA as sole carbon and nitrogen source (139).
GABA metabolism to succinate involves transport, transaminase-dependent deamination, and oxidation. GabP specifically transports GABA, and gabP mutants cannot use GABA as a nitrogen source (237, 350). GabP is a member of the amine-polyamine-choline superfamily of transporters and has been used as a model to study their structure and function (e.g., see references 255 and 256). The gabT gene specifies a GABA-glutamate transaminase, and gabT mutants cannot utilize GABA as a nitrogen source (315, 456). GabT has been partially purified and characterized (25, 459). The gabD gene specifies an NADP-dependent succinic semialdehyde dehydrogenase. The isolation of GABA nonutilizers has not resulted in mutants lacking dehydrogenase activity (315). A gabD mutant has not been constructed, but loss of GabD may have no phenotype because of the existence of multiple isozymes. E. coli B and related bacteria have another dehydrogenase that can use either NAD or NADP, although NAD is preferred (114, 115). GABA and succinic semialdehyde appear to induce this enzyme (114, 115). These results are not consistent with those reported for E. coli K-12, in which GABA induces only GabD and which is strictly NADP-dependent. The induction by succinic semialdehyde suggests that this unidentified NAD/NADP-dependent dehydrogenase is not the putrescine-inducible dehydrogenase.
The operon structure of the gab genes is complex and depends on the environment. The gene sequence of gabD-gabT-gabP was correctly established only after the entire region had been sequenced (26, 350). Nitrogen limitation induces the permease, transaminase, and dehydrogenase in E. coli and K. aerogenes (114, 116, 139, 237, 456) and coordinately elevates their transcripts (587). An insertion in gabT and deletion of gabC/csiR both dramatically increase GabD synthesis, which strongly suggests that the insertion is polar on gabC/csiR and that the gab operon contains gabC/csiR (456). GABA does not induce gab operon expression (456); however, Nac activates expression during nitrogen-limited growth, and the Nac-dependent promoter has been identified (456). GabC/CsiR represses the operon, but the factor that binds this repressor, if any, is not known (456). Curiously, loss of GabC/CsiR does not affect expression of an operon fusion of the gabD control region to lacZ (456). Instead, it appears that GabC/CsiR controls a σS-dependent promoter of a larger csiD-ygaF-gabDTPC operon (316). An insertion in ygaF has no effect on utilization of GABA as a nitrogen source, which implies that gabDTPC from the promoter preceding gabDis sufficient for GABA catabolism during nitrogen-limited growth (456). A σS-dependent promoter preceding gabD has also been identified (316).
Several environmental factors regulate expression of the gab operon. It is frequently not known if these factors activate transcription from a four- or six-gene operon. Nitrogen limitation and entry into stationary phase have been discussed. The gabP gene and presumably the gab operon were identified as one of the ten most strongly expressed σS-dependent genes for cells grown in nutrient broth (447). Conditioned medium from cells grown in nutrient broth also induces the gab operon (23). Indole is one of two active factors in conditioned medium and accounts for about 25% of the activation (552). Similar regulation was observed for the astCADBE operon (552). The other inducing signal has not been determined, although it might be a nucleoside (235). Finally, an alkaline environment appears to induce expression of gabT and probably other products of one of the gab operons (497). This regulation may be of physiological significance, since indole is the product of tryptophanase, which is also induced in alkaline media (583).
It seems likely that the GABA catabolic pathway has more than one physiological function. The nonspecific induction of gabDTPC operon expression and the number of enzymes of polyamine metabolism induced by nitrogen limitation led to the proposal that the products of the gab operon contribute to polyamine homeostasis (456). Similarly, the induction of the six-gene csiD-ygaF-gabDTPC operon by a variety of stresses led to the proposal that its products are important during stress, perhaps also by adjusting polyamine pools (316). GabT and probably other products of the gab operon are also induced by growth in an alkaline environment (497). The role of this pathway in cytoplasmic acidification is discussed at the end of the amino acid decarboxylase section.
It appears that two pathways contribute to ornithine degradation. An E. coli astC mutant grows twice as slow as a wild-type strain with ornithine as the nitrogen source (131, 454). In Pseudomonas aeruginosa, AstA can succinylate ornithine, and enzymes of the AST pathway of arginine catabolism can further degrade succinylornithine (544). This does not appear to be the case for E. coli since AstC appears to be the only enzyme of the AST pathway required for ornithine catabolism (454). Because AstC has ornithine transaminase activity, it was proposed that AstC deaminates ornithine to glutamic semialdehyde, which is an intermediate in proline catabolism (454). Loss of ornithine decarboxylase (SpeC) also appears to impair growth with ornithine as a nitrogen source, which suggests that putrescine is an intermediate in ornithine catabolism (471). Two previously described arginine transport systems, the argT (LAO) and ArgBP-II/AO (ArtI) systems, also transport ornithine. In addition, potE transports ornithine (215, 216, 352). Nitrogen limitation induces AstC and ArgT, but not PotE or SpeC (268, 454, 470, 587).
Some strains of E. coli can use tryptophan inefficiently as a sole carbon source (349, 486, 581). E. coli cannot use tryptophan as a sole nitrogen source in glucose-containing medium because of catabolite repression control of tryptophanase synthesis, but presumably it can use tryptophan with other non-catabolite-repressing carbon sources. Serovar Typhimurium cannot use the aromatic amino acids as carbon or nitrogen sources (175). K. aerogenes can use tryptophan, phenylalanine, or tyrosine as the sole nitrogen source (371).
The only enzyme of aromatic amino acid catabolism in E. coli appears to be tryptophanase. Tryptophanase catalyzes the reversible degradation of tryptophan to indole, pyruvate, and ammonia. Tryptophanase is present in E. coli and some Klebsiella species (190, 299). Most clinically isolated Klebsiella species are either Klebsiella pneumoniae or Klebsiella oxytoca. The latter produces indole from tryptophan, but the former does not. However, indole-positive and indole-negative strains can be isolated from a single individual, and it was therefore concluded that the strains were clonal and that generation of indole may be insufficient to distinguish Klebsiella species (299). Salmonella species do not contain tryptophanase (149, 486).
Tryptophanase activity requires pyridoxal-5'-phosphate and a monovalent cation, either K+ or NH4+ (486). The enzyme is a homotetramer, has a slightly alkaline pH optimum, and has a Km for tryptophan of 0.3 mM (486). In the reverse direction, the Kms for indole, pyruvate, and ammonia are 0.02 mM, 70 mM, and 345 mM, respectively (486). Tryptophanase also accepts cysteine (Km, 11 mM) and serine (Km, 160 mM) (486) as substrates and degrades cysteine in vivo (22). The pyridoxal-5'-phosphate cofactor is catalytically active at low pH and inactive at high pH; however, tryptophan converts the cofactor to the catalytically active form (218). The structure of tryptophanase has been determined (221), and details of the catalytic mechanism have been analyzed (380 and references therein). Tryptophanase has 43% amino acid identity with tyrosine phenol-lyase, which degrades tyrosine to phenol, pyruvate, and ammonia. The latter is found in several Enterobacteriaceae, but not in E. coli or serovar Typhimurium (304).
Tryptophanase is encoded by the tnaA gene and is in a tnaAB operon that also specifies a low-affinity permease (102, 120). Expression is controlled by catabolite repression and tryptophan-specific induction. Tryptophanase synthesis is very sensitive to glucose repression, which was first observed in 1936 (191). This repression accounts for the inability of E. coli to use tryptophan as sole nitrogen source in glucose-containing medium. Pyruvate represses almost as well as glucose (137). This effect of pyruvate may be mediated by cyclic AMP. However, there are other documented mechanisms by which pyruvate can negatively affect protein activity (described in the section on the biodegradative threonine deaminase). Cyclic AMP and CRP induce tryptophanase synthesis (375), and they appear to be the only factors that mediate catabolite repression (unpublished results cited in reference 304). They are also required for transcription in vitro (103). Certain aspects of catabolite repression, however, remain unexplained (44); for example, cyclic AMP induces expression even after the addition of several RNA synthesis inhibitors (375). It was originally stated that translation might be required for transcription (375), which was prescient considering the unusual attenuation mechanism that mediates tryptophan-specific control. Transcription is terminated before the tnaA structural gene without tryptophan, but continues into tnaA with tryptophan (505). Induction requires a 24-residue leader peptide, TnaC (506), and tRNAtrp-dependent translation of codon 12 (156). Furthermore, induction requires tryptophan-12 in the nascent TnaC in the peptide exit tunnel of the ribosome (159, 260, 262). This inhibits release factor-mediated cleavage of the TnaC-peptidyl-tRNA in the translating ribosome (157, 158, 160, 582). The ribosome stalls, which blocks the binding sites for the termination factor ρ (158, 160, 261, 504, 505, 580). Coupling of transcription with translation is achieved with a transcriptional pause site at the end of the tnaC coding region (161).
Catabolite repression and tryptophan-specific induction suggest that tryptophanase has a catabolic function, specifically that it provides carbon and energy. However, McFall and Newman (304) questioned whether such a catabolic function was consistent with the distribution of tryptophanase in the Enterobacteriaceae. Such a catabolic function predicts that tryptophanase-containing organisms would have a competitive advantage in an environment with tryptophan, but this is not the case (45). Other factors control tryptophanase expression, which suggests that it has further functions. Tryptophanase is induced by depletion of heme (421) or phosphatidylglycerol (359), aerobic or anaerobic growth in alkaline broth (39, 497, 583), anaerobic growth with the electron acceptor trimethylamine N-oxide (43), cysteine (22), and the product indole (23, 552). Tryptophanase is repressed by acetate (257). To account for many of these effects, it has been proposed that tryptophanase alleviates alkaline stress (39, 43, 497, 583). This hypothesis proposes that tryptophan degradation produces NH3 (which is excreted) and pyruvate (which generates such acids as acetic acid), which acidifies the cytoplasm. This hypothesis receives strong support from the observation that a tryptophanase mutant is defective in survival in an alkaline environment (43). It may also explain the apparent induction by cysteine (22), which may alkalinize the medium. Survival in an alkaline environment is consistent with the massive overproduction of tryptophanase; tryptophanase can accumulate to as much as 10% of the total cytoplasmic protein (55, 486) and can be the major protein in some two-dimensional gels of cell extracts (39, 421). An additional function of tryptophanase may be to synthesize the intercellular signaling molecule, indole, which regulates gene expression and biofilm formation (111, 298, 552).
K. aerogenesmutants unable to utilize tryptophan as a nitrogen source lack a multispecific aromatic amino acid transaminase (371). The purified dimeric transaminase can use phenylalanine (Km, 0.08 mM), tryptophan (Km, 0.6 mM), and probably tyrosine as amino group donors (370). Transaminase synthesis is constitutive, but synthesis of a tryptophan transport system is under Ntr control (371). The genes for the transaminase and the transport system have not been identified. E. coli does not contain the catabolic aromatic amino acid transaminase (L. Reitzer, unpublished data).
Six transport systems have been identified for the aromatic amino acids (263, 382). They are present in E. coli and serovar Typhimurium, and their functions have been recently reviewed (382). Tryptophan is transported by a general aromatic amino acid transporter, AroP, and by two tryptophan-specific systems, Mtr and TnaB (581). Mtr also transports indole (581). TrpR, TyrR, or both control their synthesis (382). The tnaAB operon and the factors that control this operon were described above. Growth with tryptophan as a carbon source requires TnaB (581), which has 40 times the capacity of Mtr (382). Expression of the mtr gene is repressed by tryptophan (via TrpR), and in the absence of tryptophan expression is activated by phenylalanine or tyrosine (via TyrR) (382). Expression of aroP is repressed by each of the aromatic amino acids (via TyrR) (382). Tyrosine is transported by AroP and the tyrosine-specific transport protein, TyrP. Expression of the gene specifying TyrP is repressed by tyrosine (via TyrR) and activated by phenylalanine (via TyrR) (382). Phenylalanine is transported by AroP, the phenylalanine-specific PheP, and the LIV-I/LS system of branched-chain-amino-acid biosynthesis (263, 382). Both binding proteins of the latter system contribute to phenylalanine transport (263). All the systems, except for the LIV-I/LS system, are energized by the proton motive force (382). With the exception of tnaB, the capacities or regulation of the other systems are not obviously consistent with a catabolic function. Instead, they appear to provide amino acids for protein synthesis and to ensure a proper balance of the aromatic amino acids.
E. colican utilize aspartate as either a carbon or a nitrogen source and glutamate as a nitrogen source. Wild-type E. coli cannot utilize glutamate as a carbon source, although mutants that can utilize glutamate as a carbon source have been isolated (182, 183, 545, 546). Serovar Typhimurium can degrade either amino acid as a nitrogen source, but neither as a carbon source (175). A major component of the catabolism of these amino acids is deamination. As discussed above, E. coli grown aerobically requires about 80 mmol of glucose carbon per gram of cells (dry weight). With glutamate or aspartate as a carbon source, growth requires 16 mmol of glutamate or 20 mmol of aspartate. About 15 mmol of glutamate or 18 mmol of aspartate must be deaminated, which takes into consideration the direct assimilation of the entire carbon-nitrogen skeleton into other compounds. The nitrogen requirement is substantially lower. One gram of bacteria requires about 10 mmol of nitrogen, with 8.2 mmol for nitrogen derived from glutamate (which does not require deamination of glutamate or aspartate), and 2.3 mmol for glutamine synthesis (which requires ammonia-generating deamination). In other words, the quantitative significance of the ammonia-generating deaminations is at least six times greater for carbon source utilization than nitrogen source utilization. The pathways for glutamate or aspartate catabolism might depend on whether they are used as a nitrogen or a carbon source. Transport and possible catabolic pathways are summarized in Fig. 6.
Catabolism of glutamate as a carbon source has been more extensively studied than that of aspartate, even though wild-type E. coli can utilize the latter but not the former. The starting point for these studies has been isolation of mutants that utilize glutamate as a carbon source (182, 183, 545, 546). These mutants generally have elevated glutamate transport (183, 297, 545). It was subsequently shown that such growth correlates with elevated transport by either the Na+-dependent GltS (GltC in earlier publications) or the H+-dependent GltP transport system. (The characteristics of the four glutamate and six aspartate transport systems are more thoroughly described in Chapter Biosynthesis of Glutamate, Aspartate, Asparagine, l-Alanine, and d-Alanine.) The glutamate-utilizing mutants had normal levels of aspartase (AspA) and aspartate transaminase (AspC), but reduced glutamate dehydrogenase and glutamate decarboxylase (182, 545, 546). Glutamate is converted to γ-aminobutyrate in a wild-type strain, and to α-ketoglutarate in a glutamate-utilizing mutant (545). This and an observation by Jung and Kim (236) appear to be the only evidence that glutamate decarboxylase has a function during growth, in addition to a function during acid survival and stationary phase. Although never explained, perhaps glutamate decarboxylase diverts glutamate away from deaminating reactions. The glutamate-utilizing mutants were further mutagenized to determine the pathway of glutamate catabolism. These mutants were defective in glutamate transport, aspartase, or aspartate transaminase (297). Marcus and Halpern proposed that the transaminase AspC aminated oxaloacetate and that aspartase degraded the resulting aspartate to ammonia and fumarate (297). It was subsequently shown that transduction from a wild-type strain to the aspartase mutant could not be used to isolate a glutamate utilizer in a single step, which implies that the mutant has at least two unlinked lesions that prevent aspartate catabolism (492). Aspartase also appears to be required for utilization of aspartate as a carbon source (495).
Attempts to genetically analyze the use of glutamate or aspartate as the sole nitrogen source have not been successful. Two genetic screens resulted in isolation of asnB and nit/nadE mutants, which have pleiotropic defects in nitrogen source utilization (50, 412, 455). An alternative strategy to study aspartate catabolism was an NMR analysis of metabolites derived from 15N-aspartate, which showed rapid labeling of glutamate, alanine, and putrescine (167). The possibilities that alanine or arginine were intermediates in aspartate degradation were specifically examined and disproven (454, 555). Nonetheless, evidence suggests that aspartase contributes to aspartate degradation. Mutants that showed improved growth with aspartate as a nitrogen source had elevated levels of aspartase (353). Furthermore, the presence of aspartase correlates with improved growth of Yersinia spp. (118).
It is possible that more than one enzyme contributes to aspartate catabolism. An obvious candidate for an aspartate or glutamate catabolic enzyme is glutamate dehydrogenase. However, nitrogen limitation represses its synthesis, which suggests an anabolic function, and gdhA mutants grow normally with glutamate as nitrogen source (see Chapter Biosynthesis of Glutamate, Aspartate, Asparagine, l-Alanine, and d-Alanine). Glutamate decarboxylase cannot degrade glutamate, since ammonia is not generated by the decarboxylation or subsequent degradation of GABA. Several isolated observations have been made concerning catabolism of aspartate as a nitrogen source. An unidentified gene near ansA may be required for aspartate degradation (495). Elevated expression of the arginine catabolic astCADBE operon stimulated growth with aspartate as a nitrogen source, but loss of this operon did not impair growth (454). Kay proposed that ast, which has not been localized, specifies a major transport system for aspartate as a nitrogen source (248). The ast mutant is resistant to β-aspartyl hydroxamate, and a subsequent study suggested that such resistance might result from loss of two aspartate transport systems (446). Overexpression of the GltP transport system enhances growth with aspartate as a nitrogen source (446), but does not necessarily imply a role for GltP in a wild-type strain. Aspartate as sole nitrogen source induces the Ntr response (413), yet ntrC mutants grow normally with aspartate (368), which suggests that no essential component of aspartate catabolism is regulated by the Ntr system. The only known component of aspartate/glutamate metabolism under Ntr control is the GltIII (gltIJKL) transport system (587). The intriguing observation that aspartase binds to NtrB suggests a potential regulatory role for aspartase in the control of nitrogen source utilization (436).
Aspartase catalyzes the reversible conversion of aspartate to fumarate and ammonia. The enzymology of aspartase has been reviewed by Viola (549). Aspartase from E. coli is a homotetramer from all reported sources, and the subunit Mr for the E. coli K-12 enzyme is 54,064, although other sizes have been reported (174, 429, 516, 523, 569). Partial proteolysis of aspartase increases its activity, which could account for the diversity of sizes reported (226, 549). The enzyme is very specific for aspartate (549). The apparent Km is in the low millimolar range and depends on such factors as pH, metal ions, and allosteric activators (549). At neutral or acidic pHs, the kinetics are not unusual. At alkaline pHs, the kinetics are nonlinear, and a divalent metal ion and an activator are required for maximal enzyme activity. The activator can be L-aspartate and a variety of other compounds that do not act as substrates, including D-aspartate (238, 549). One metal ion binds per subunit, and the activator site is not near the active site (127). Karsten and Viola have proposed a pH-dependent equilibrium between two forms (239). This could explain occasional reports of two forms of aspartase, including a partial membrane localization (143, 532). However, it has not been possible to fractionate the two forms or to detect a pH-dependent conformational change in the enzyme (238, 429, 516). The structure of aspartase has been determined, and active site residues have been identified (227, 474, 549).
The monocistronic aspA gene specifies aspartase, and a transcription start site has been identified (150). Two environmental factors affect aspA expression. First, the aerobic repression of aspartase formation or aspA expression by glucose in minimal medium or nutrient broth has been known since 1938 and subsequently confirmed (124, 143, 182, 568). Mutants that are insensitive to catabolite repression have elevated aspartase activity (353). These results suggest activation by cyclic AMP and CRP, but this has not been directly tested. This regulation may prevent the participation of aspartase in the catabolism of aspartate as a nitrogen source. Second, the FNR protein during anaerobic growth activates aspA expression (233, 568). FNR-independent expression appears to account for about 25% of the maximal expression level (233, 495). FNR-dependent regulation suggests that an important function of aspartase is generation of fumarate, an anaerobic electron acceptor (233, 493). Potential CRP and FNR sites have been located but have not been shown to be functional (568). In addition to these factors, glutamate induces aspartase synthesis in a mutant that can utilize glutamate as a carbon source, but not in a wild-type strain (545, 546). Nitrogen limitation has been reported to severely repress aspartase (116), although a microarray analysis of Ntr genes did not confirm this result (587). A serovar Typhimurium mutant with elevated glutamine synthetase activity that grew faster with aspartate as a nitrogen source had wild-type aspartase activity, which further indicates lack of Ntr control (142).
E. coli, K. aerogenes, and serovar Typhimurium can utilize asparagine as the sole nitrogen source. E. coli can also use asparagine as the sole source of carbon (495). These bacteria contain a high-affinity periplasmic asparaginase and a low-affinity cytoplasmic asparaginase. Therefore, there are two distinct pathways of asparagine utilization: asparagine transport, followed by asparagine hydrolysis in the cytoplasm, or asparagine hydrolysis in the periplasm, followed by ammonia or aspartate transport (Fig. 6). E. coli mutants unable to use asparagine as a nitrogen source lack either the cytoplasmic asparaginase (AnsA) or a low-affinity asparagine transport system (AnsP or Asu) (81, 104, 561). Loss of the cytoplasmic asparaginase also blocks utilization of asparagine as a carbon source (495). These results imply that E. coli transports asparagine before degradation, regardless of whether it is being used as sole carbon or nitrogen source. In contrast, a K. aerogenes mutant lacking the periplasmic enzyme grows significantly slower than a wild-type strain, which underlines the importance of both pathways (414).
The cytoplasmic asparaginase, also called asparaginase I, is the product of ansA. The dimeric E. coli enzyme has a Km for asparagine of 3.5 mM (232, 561). ansA is part of an ansA-ydjB operon; the second gene specifies a protein of unknown function (232). Expression is considered constitutive (517) and does not respond to nitrogen limitation (587). However, there are hints of regulation, since enhanced synthesis of asparaginase II correlated with decreased asparaginase I levels and the extremely high expression of ansA from a plasmid suggested titration of a repressor (495, 517).
The periplasmic asparaginase, also called asparaginase II, is the product of ansB. Asparaginase I and II are not homologous (229, 232). Asparaginase II has been purified from E. coli and K. aerogenes (205, 414). The E. coli enzyme has a Km for asparagine of about 10 μM (205). The factors that control synthesis of the periplasmic enzyme in E. coli and serovar Typhimurium imply that its primary function is generation of fumarate as an electron acceptor during anaerobic respiration (233, 493). In E. coli, maximal expression requires anaerobic growth, the lack of glucose, and a mixture of amino acids (66, 430). CRP mediates the glucose effect, FNR mediates the response to anaerobic conditions, and a novel regulator, Arr, mediates the response to amino acids ( 170, 229, 230, 233, 430). In contrast, FNR does not control ansB in serovar Typhimurium (231). In K. aerogenes, nitrogen limitation, not anaerobiosis, induces synthesis of the periplasmic asparaginase (414). The induction does not require asparagine (414).
E. coligrown in ammonia-containing (nitrogen-rich) medium has two kinetically distinct components for asparagine transport (562). Asparagine represses the high-affinity system (562). The ansP gene specifies the low-affinity system (228), which is almost certainly the previously described asu locus (81). Although this protein is required for utilization of asparagine as a nitrogen source, nitrogen limitation has no effect on its synthesis (587).
Wild-type E. coli grows very slowly with glutamine as sole carbon source, although fast-growing mutants can be isolated (300). Glutamine is a signal of nitrogen sufficiency and, not surprisingly, supports reasonably rapid growth as sole nitrogen source. Nonetheless, glutamine as a nitrogen source induces the Ntr response, probably a result of very rapid catabolism.
More is known about glutamine transport than about the enzymes of glutamine catabolism. E. coli and serovar Typhimurium contain at least two glutamine transport systems (Fig. 6) (36, 557, 560). The high-affinity system, specified by the glnHPQ operon, is an ABC cassette system (300, 355). Glutamine is the only naturally occurring amino acid that binds the periplasmic binding protein (557, 560). A mutant that can more efficiently utilize glutamine as a carbon source has 2.5 to 6 times more of the glutamine-binding protein than a wild-type strain (300, 557). This mutant can utilize glutamate as a carbon source, and derivatives of this mutant with reduced glutamine transport cannot utilize glutamate as a carbon source (300). The structure of the binding protein from E. coli has been determined (211). Nitrogen limitation induces the high-affinity transport system in E. coli, serovar Typhimurium, and K. aerogenes (36, 268, 560, 587) (L. Reitzer, unpublished data). Transcription of the glnHPQ operon can initiate from two promoters, an Ntr-regulated promoter and a constitutive promoter (354). NtrC and integration host factor are required for expression from the Ntr-regulated promoter (83). In addition, Fis moderately represses glnHPQ expression (576). A low-affinity system is detectable either in a mutant lacking the high-affinity system or in cells with repressed high-affinity transport activity (557, 560). The metD gene, which specifies a high-affinity methionine transporter, has been proposed to specify the low-affinity glutamine transporter (P. D. Ayling, unpublished evidence cited in reference 171). Glutamate inhibits the low-affinity glutamine system, which suggests that a glutamate carrier might also be a low-affinity glutamine carrier (557).
As mentioned above, glutamine is the signal of nitrogen sufficiency, but it is a growth-limiting nitrogen source. Glutamine as a nitrogen source supports biphasic growth, and cells become nitrogen limited only during the slower growth phase (20). The basis for this unusual growth is not understood, possibly because the enzymes of glutamine catabolism are not known with certainty. Mutants unable to utilize glutamine as a nitrogen source, and thus lacking all glutamine-degrading enzymes, have not been isolated. Currently, only possible glutamine-catabolizing enzymes can be considered. For example, mutants deficient in glutamate synthase (GOGAT) grow slowly with glutamine as the sole nitrogen source, which suggests that it is an important enzyme of glutamine catabolism (165). E. coli also contains several enzymes with glutaminase activity: glutaminase A, glutaminase B, asparaginase II (which should not be expressed during exponential nitrogen-limited growth), and a side reaction of amidotransferases. Glutaminase A is a homotetrameric protein with subunits of about 28,000 (194). It is active only at acidic pH (pH 5) (193). It is induced by energy depletion, and this induction is prevented by cyclic AMP and CRP (396, 398). Glutaminase B may also be homotetrameric; its mass is about 90,000 (395, 397). Glutaminase B is apparently constitutive (396). It is active at neutral pH, activated by AMP and divalent cations, and inhibited by ATP and ADP (397, 399). The regulatory factors that control glutamine synthetase and glutaminase B may prevent futile cycling (399). Consistent with this conclusion is the observation that nitrogen limitation does not induce potential glutamine-degrading enzymes. Nitrogen limitation represses GOGAT (see Chapter Biosynthesis of Glutamate, Aspartate, Asparagin, l-Alanine, and d-Alanine) and glutaminase A, but has no effect on the allosterically regulated glutaminase B (396). The genes for the glutaminases have not been identified, and mutants deficient in either glutaminase have yet to be isolated. The physiological functions of the glutaminases have not been established.
Serovar Typhimurium can degrade cysteine (or cystine) as sole carbon or nitrogen source, and K. aerogenes can utilize cysteine as a nitrogen source (175, 538). E. coli can very slowly utilize cysteine as sole nitrogen source (W. Gray and L. Reitzer, unpublished observation). Transport systems and catabolic enzymes are summarized in Fig. 7.
Cysteine (or cystine) inhibits the growth of many strains of E. coli and serovar Typhimurium (86, 420, 428, 488). The inhibition is bacteriostatic and potentially bacteriocidal. The bacteriostatic effect results from inhibition of various enzymes. Cysteine induces transient amino acid starvation (488), but paradoxically decreases guanosine tetraphosphate levels (105). Methionine, isoleucine, threonine, or homoserine overcomes cysteine toxicity (105,192, 428). Cysteine inhibits threonine deaminase and homoserine dehydrogenase I, which would account for stimulation by threonine and isoleucine (98, 184, 192). MetC, cystathionine β-lyase, has cysteine desulfhydrase activity (see below), and presumably cysteine inhibits lyase activity, which could account for the stimulatory effect of methionine. Cysteine also inhibits L-serine deaminase (2, 82). Because alleviation of toxicity by amino acid supplementation is only partial, other cysteine-sensitive targets probably exist. Cysteine and serine appear to inhibit similar enzymes.
The bacteriocidal effects of cysteine are mediated by sulfide production. Sulfide inhibits the primary enzyme of cysteine catabolism, cysteine desulfhydrase, at least in serovar Typhimurium, which should minimize toxicity (85, 86, 173, 264). However, cysteine and sulfide, in particular, potentiate hydrogen peroxide killing (33, 34). This killing was abolished by the iron-chelating agent 2,2'-bipyridyl (34). Iron sulfide may stimulate the Fenton reaction, producing hydroxyl radicals and resulting in DNA breakage (33, 34). Formation of iron sulfide may account for the observation that a mutant deficient in cysteine export (resulting in intracellular cysteine accumulation) becomes iron limited (discussed below).
The primary enzyme of cysteine catabolism is cysteine desulfhydrase, which degrades cysteine to pyruvate, ammonia, and H2S. Cysteine catabolism is readily detectable in E. coli, even in complex medium, because of H2S production. E. coli has been known to produce H2S from cysteine since at least 1939, and cysteine desulfhydrase has been assayed from crude extracts (references 11 and 420 and references cited therein). Furthermore, cysteine catabolism limits both the growth of cysteine auxotrophs (420) and the overproduction of cysteine from genetically altered E. coli (330).
Four different enzymes have cysteine desulfhydrase activity: tryptophanase (TnaA); cystathionine β-lyase (MetC); tryptophan synthase; and a dedicated cysteine desulfhydrase (at least in serovar Typhimurium) (84, 85, 92, 119, 264, 265, 427, 486). The cysteine biosynthetic enzyme, CysK or O-acetylserine sulfhydrylase A, might catalyze a reverse reaction that would be equivalent to the activity of cysteine desulfhydrase (497). However, the reverse reaction is only about 1% of the biosynthetic reaction, and strong arguments have been presented to indicate that this reverse reaction does not occur in vivo (129). All these enzymes require pyridoxal-5'-phosphate. In E. coli, TnaA and MetC have cysteine desulfhydrase activity, and a double mutant lacking both enzymes still retains 30% of the total activity (22). Some 40 years ago, coregulation (repression by methionine) of cystathionine β-lyase (MetC) and cysteine desulfhydrase activities suggested that the same enzyme catalyzed both reactions in E. coli (427). Cysteine induces cysteine desulfhydrase activity (173), and this was probably due to TnaA, which is also induced by cysteine and which becomes the predominant cysteine desulfhydrase in the presence of cysteine (22). The mechanism of induction is not known.
Serovar Typhimurium contains a cysteine-inducible cysteine desulfhydrase, which was clearly not cystathionine β-lyase (86, 173). The purified enzyme has essentially no tryptophanase, cystathionine β-lyase, or tryptophan synthase activity, which suggests that it is a dedicated cysteine desulfhydrase (265). The kinetics are sigmoidal with respect to cysteine concentration (approximate Km of 0.2 mM) (84, 86, 264). This may prevent cysteine depletion when the desulfhydrase is optimally expressed (86). The purified enzyme has a Ki for sulfide of about 10 μM, which probably prevents toxic sulfide accumulation (85, 264). Nonsubstrate cysteine analogs (e.g., serine, methionine, and homocysteine) stimulate activity, and it was suggested that these compounds may bind to a cysteine-specific regulatory site (85). Such regulation amounts to an elaborate feedback mechanism, since excess homocysteine and methionine will presumably lower the concentration of cysteine, which is required for their synthesis. Two groups isolated cysteine desulfhydrase from the same strain, but reported slightly different sizes for the enzyme (Mr 229,000 versus 184,000) (85, 265). The protein appears to be a hexamer (265). Its gene has not been identified. Curiously, the ratio of the two products, sulfide and pyruvate, changes on purification of the enzyme (84, 85, 264, 265). This appears to result from the formation of a conjugate between pyruvate and cysteine, 2-methyl-2,4-thiazolidinedicarboxylic acid (264). The expected product ratio for the reaction is restored by a protein that had been separated from cysteine desulfhydrase during purification. This protein was not further characterized (84, 264).
D-Cysteine desulfhydrase is widespread in the family Enterobacteriaceae, including E. coli, where it was first detected in 1954 (327, 443, 491). The purified enzyme is a homodimer with a subunit mass of 35,153 Da (327, 491). It contains pyridoxal-5'-phosphate (327, 491) and stoichiometrically generates pyruvate, NH3, and H2S (327). The Kms for D-cysteine and D-cystine are about 0.2 mM and 0.3 mM, respectively (327, 491). L-Cysteine is an inhibitor (Ki, 0.5 mM) (327). Enzyme activity is not induced by the L- or D-isomers of cysteine, cystine, serine, or methionine (327, 491). Instead, synthesis of the enzyme is induced by sulfur starvation, suggesting that the enzyme scavenges sulfur (491). The gene encoding D-cysteine desulfhydrase is yedO(491). D-Cysteine, like L-cysteine, inhibits growth (491). This inhibition is overcome by aspartate or the branched-chain amino acids (491). One site of inhibition is threonine deaminase, but other enzymes are probably also inhibited (491). A ΔyedO mutant is viable but retains only 3 to 10% D-cysteine desulfhydrase activity, which presumably results from D-amino acid dehydrogenase (491). Loss of the desulfhydrase results in D-cysteine hypersensitivity but has no effect on L-cysteine sensitivity (491). The yedO gene may be part of an operon that also contains a transport system for L- or D-cystine (491).
Transport studies with whole cells suggest three kinetically distinct cystine transport systems, CTS-I, CTS-II, and CTS-III, in serovar Typhimurium (24). CTS-I accounts for more than 90% of cystine transport (at least at a certain cystine concentration) (24). Osmotic shock reduces transport by 34%, which suggests an effect on CTS-I (24). Sulfate limitation induces CTS-I but has no effect on CTS-II (24). Synthesis of CTS-I requires CysB, and a cysB mutant is unable to induce cysteine desulfhydrase (24).
Whole-cell transport studies with E. coli suggest two cystine transport systems (32, 273). One system transports both cystine and meso-diaminopimelate (transport Km, 0.3 μM), and the second is more specific for cysteine and has a higher affinity (transport Km, 0.02 μM) (32, 273). Growth in nutrient broth represses the less specific and induces the more specific transport system (32). Osmotic shock reduced the activity of the former, which suggests that it is the equivalent of CST-I (32). A binding protein from the shock fluid was purified and characterized by two groups (cystine Kd, 0.01 to 0.03 μM; diaminopimelate Ki, 17 μM) (32, 57). From the amino acid composition and amino-terminal amino acid sequence (32, 57), it can be deduced that fliY codes for the binding protein (B. L. Schneider, unpublished observation). The membrane components for the cysteine-binding protein have not been determined. fliY is adjacent to yedO, which codes for D-cysteine desulfhydrase (see above). The fliY gene is a member of the fliAZY operon and fliA specifies σF, and fliZ codes for a protein of unknown function (326). Sigma factor σF is required for expression of class III genes of the motility regulon. A ΔfliZY strain is motile, although it forms smaller swarms on semisolid agar and more than 99% of cells are nonmotile in liquid medium. Complex genetic results are consistent with the possibility that FliZ and FliY might negatively regulate fliA (326). The fliA gene has two promoters, one of which is a σ70-dependent promoter that requires FlhD, and the other is σF dependent and does not require an activator in vitro. Loss of FlhD not only eliminates transcription from the former but, unexpectedly, reduces transcription by 80% from the latter (326). Numerous factors control FlhD synthesis, which are beyond the scope of this review.
CST-II, or its equivalent, has not been well characterized. Cysteine transport has been characterized from E. coli membrane vesicles, that is, binding protein-independent transport (282). Cysteine transport is not inhibited by any of the protein amino acids, hydroxyproline, ornithine, or D-serine (282), and this activity probably represents the cysteine-specific CST-II. Such transport is driven by the proton motive force and has a transport Km of 38 μM (282). Curiously, the cysteine transporter has the lowest Km of any vesicular transport system, but the highest capacity (Vmax) (282). Isoleucine, leucine, and valine stimulate cysteine export from membrane vesicles (282). Leucine, valine, or alanine overcomes a cysteine toxicity (33), which is at least consistent with the possibility that this export is a physiological process, perhaps mediated by a cysteine export system (discussed below).
E. colipossesses two cysteine export systems. One system consists of CydD and CydC (383) and there is abundant literature on mutants lacking these proteins. Both proteins are required for cytochrome bd activity (147, 386, 388). Cytochrome bd is a membrane-bound terminal oxidase that is induced by low oxygen. The loss of CydC or CydD results in a surprising variety of phenotypes, including loss of periplasmic b- and c-type cytochromes, loss of motility, sensitivity to high temperature, peroxide, zinc, and azide, and inability to exit from the stationary phase (reference 383 and references therein). The mutant also excretes enterochelin, which suggests iron limitation (87). Overexpression of cytochrome bd, or low-molecular-weight reducing agents, including a nonpermeant reducing agent, reverses these defects (154). The latter finding suggested an alteration in the periplasmic redox state, which was confirmed by examining the availability of sulfhydryl groups in periplasmic proteins (155). CydC and CydD are components of a heterodimeric ABC transport system (386) that exports cysteine to the periplasm (383). Mutants lacking these proteins accumulate cysteine more efficiently, have elevated intracellular cysteine, and are therefore hypersensitive to cysteine toxicity (105, 383). It was concluded that cysteine contributes to redox homeostasis in the periplasm, and loss of the export system results in a more oxidized periplasmic environment (383). The cydD mutation is allelic to htrD. However, HtrD has been reported to be a 17-kDa protein, whereas CydD is a 65-kDa protein (106, 387). Comparing the genome sequences of strains W3110 and MG1655 indicates that the cydD gene from W3110 has a guanosine nucleotide deletion at codon 101, which results in a truncated protein product relative to that encoded in MG1655. The cydDC operon is expressed aerobically, or anaerobically with alternative electron acceptors (88).
The ydeD gene specifies a second potential cysteine export system (97). When expressed at a high level, YdeD exports cysteine in the form of 2-methyl-2,4-thiazolidinedicarboxylic acid (the reaction product of the cysteine desulfhydrase reaction), and intermediates of the cysteine biosynthetic pathway, such as O-acetyl-L-serine (97). Such cells require reduced sulfur compounds for growth in minimal medium (97). A strain with a deletion of ydeD was initially reported to have no observable phenotype (97), although such a mutant is hypersensitive to cysteine, and a ydeD cydD double mutant is even more sensitive than either single mutant (383).
There appears to be only one reference to utilization of methionine as a nitrogen source, and it suggests that E. coli can slowly degrade methionine as sole nitrogen source (368). However, this observation has not been verified and possible enzymes of methionine catabolism in E. coli and related organisms have not been studied. Methionine can be salvaged as a source of sulfur/cysteine in K. aerogenes, but not in E. coli or serovar Typhimurium (466). Several methionine salvage pathways exist in bacteria (466, 467, 550). Pathways could be proposed that recycle S-adenosylhomocysteine, decarboxylated S-adenosylmethionine, and possibly homocysteine (a transsulfurylation pathway) and, at least theoretically, that might also degrade methionine. However, these would be obligatorily coupled to S-adenosylmethionine-dependent methylations or decarboxylated S-adenosylmethionine-dependent spermidine synthesis, which seem to lack the capacity to provide sufficient nitrogen. The most likely pathway for the catabolism of methionine, when used as a nitrogen source, would not be restricted by such metabolic coupling. There are two possible pathways. One is initiated by methionine γ-lyase, which produces α-ketobutyrate, ammonia, and methanethiol (550). However, K. aerogenes, which appears to have a greater capacity to degrade methionine than E. coli or serovar Typhimurium, does not contain this enzyme (466). The second pathway involves methionine transaminase, which produces 2-keto-4-methylthiobutyrate, that is, deaminated methionine, a product of the pathway that recycles decarboxylated S-adenosylmethionine (550). The tyrosine transaminase, TyrB, can catalyze the methionine transaminase reaction, at least in K. pneumoniae (197). E. coli contains another methionine transaminase, YbdL (112). It is unclear which direction the reaction occurs in vivo. A lacZ fusion to ybdL indicates stationary phase induction of expression (L. Reitzer, unpublished data). K. aerogenes also contains a methionine-inducible cystathionine-γ-lyase, which probably cleaves L-allo-cystathionine (which is generated from homocysteine) to cysteine, α-ketobutyrate, and ammonia (466). This pathway is possible, if methionine can be converted to homocysteine without obligatory coupling to the demethylation of S-adenosylmethionine.
Histidine can serve as sole source of carbon or nitrogen for K. aerogenes (292, 336, 416) (reviewed in reference 291). Serovar Typhimurium has the enzymes of histidine degradation (416). Some, but not all, serovar Typhimurium strains can utilize histidine as a nitrogen source, but not as a carbon source (49, 305). Derivatives of these strains can be isolated that can utilize histidine or urocanate (a product of histidine catabolism), as carbon or nitrogen source (49, 305). E. coli lacks the genes and enzymes for the degradation of histidine (416).
The pathway for histidine catabolism as a carbon or nitrogen source is the same (Fig. 8). The first enzyme of histidine degradation is L-histidine ammonia-lyase or histidase (HutH), which generates urocanate and ammonia. Urocanase (HutU) degrades urocanate to 4-imidazolone-5-propionate, which 4-imidazolone-5-propionate amidohydrolase (HutI) converts to N-formimino-L-glutamate. Finally, N-formimino-L-glutamate hydrolase (HutG) degrades N-formimino-L-glutamate to glutamate and formamide. Formamide is not further metabolized. The first three steps are common to all known pathways, but the fate of N-formimino-L-glutamate depends on the organism. In pseudomonads, it is degraded to ammonia and N-formyl-L-glutamate, which is cleaved to formate and glutamate. In mammalian species, N-formimino-L-glutamate generates glutamate and the formimino group is donated to tetrahydrofolate. Histidine as a nitrogen source requires ammonia generation and not glutamate degradation, although it is probably degraded. When histidine is used as a carbon source it requires glutamate degradation, and aspartase, which is induced when histidine is degraded, probably generates the nitrogen-free carbon skeleton for growth (293). The effect of loss of aspartase on growth with histidine has not been examined. Growth with histidine, or expression of histidine utilization genes, requires either oxygen or nitrate as an alternate electron acceptor and apparently a complete citric acid cycle (151, 189).
The enzymes of histidine catabolism have only been partially characterized. Urocanase (HutU) from K. aerogenes was purified threefold and has a pH optimum from 7.4 to 7.8; kinetic parameters were not assessed (415). Serovar Typhimurium 4-imidazolone-5-propionate amidohydrolase (HutI) in crude extracts has a pH optimum of 7.4, and an apparent Km for its substrate of 0.1 mM (483). K. aerogenes N-formimino-L-glutamate formiminohydrolase (HutG) was purified 40-fold and has a pH optimum of 8.5 and a Km for its substrate of 40 mM (285). Histidase (HutH) and urocanase (HutU) have been characterized from Pseudomonas putida, in which the Kms for histidine and urocanate are high, 5.5 mM and 66 mM, respectively (304).
The gene order for the hut genes in K. aerogenes and serovar Typhimurium is hutI-hutG-hutC-hutU-hutH (Fig. 8) (46, 49, 152, 291, 463, 483, 485). The hutC gene specifies a repressor. All the available evidence indicates that hutU and hutH form an operon in both organisms. Moreover, they are coordinately expressed, but not with the other hut genes, and mutations in hutU can be polar on hutH (49, 293, 305, 450, 463, 484). The products of hutI, hutG, and hutC in serovar Typhimurium are regulated coordinately, which suggests a hutIGC operon (177, 483, 484, 485). Coregulation studies have not been done in K. aerogenes. Instead, genetic evidence and sequence analysis suggest a separate promoter for each of these three genes (152, 461, 463).
Several environmental factors affect hut gene expression: induction by urocanate, catabolite repression, nitrogen limitation, and the availability of electron acceptors (Fig. 8). Changes in transcript abundance can account for the changes in the levels of hut gene products, which suggests that most regulation is transcriptional (89). Urocanate, the product of the histidase reaction, is required for induction of hut gene expression (49, 293, 305, 451). This explains why histidine does not induce expression in a histidase mutant and a urocanase mutant has high basal hut expression (49, 291, 451). The alleles hutP, hutR, and hutQ refer to sites in the hutUH regulatory region, while hutM refers to a mutation in the hutI regulatory region (291). These sites were defined by mutations that altered hut gene expression. The HutC repressor mediates urocanate induction. Serovar Typhimurium and K. aerogenes hutC mutants do not require urocanate for induction, are still subject to catabolite repression, and have increased levels of hut gene products (49, 394, 483, 484). Genetic evidence suggested HutC binds to hutM and hutP DNA (49, 291, 483). This was confirmed when partially purified HutC from serovar Typhimurium bound to DNA containing these promoters, and urocanate eliminated this binding (176, 177). Binding to hutP appears to be five times stronger than to hutM (176). HutC is unstable and has not yet been purified to homogeneity (176), but the gene has been sequenced (461). Indirect methods have identified a single HutC-binding site, which is centered at about −20 bases from the hutU transcription start site (367). HutC thus controls its own synthesis in serovar Typhimurium, and it has been suggested that a function of this autogenous regulation is to allow rapid repression of the hut operons when urocanate is depleted (291, 484).
Catabolite repression controls synthesis of the histidine utilization enzymes in K. aerogenes (292, 293, 336, 394) and serovar Typhimurium (49, 483). Cyclic AMP overcomes the catabolite repression in both serovar Typhimurium (S. Fogel and B. Magasanik, unpublished results cited in reference 485) and K. aerogenes (394). Furthermore, a K. aerogenes mutant lacking CRP could not utilize histidine as a carbon source (394). The hutU promoter region contains both a high-affinity and a low-affinity CRP-binding site, called CAP1 and CAP2, respectively (366). These sites are centered at –81.5 bp and –41.5 bp, respectively, relative to the transcription start site. The CAP1 site is essential, but the CAP2 site is required for optimal expression (366). It has been suggested that FIS might bind between the upstream site and the promoter (366). Oxygen or nitrate is required for expression (151, 189), but the possibility that FNR binds to one or more of the CRP sites and mediates this control has not been tested. There is no evidence that directly implicates cyclic AMP or CRP in control of the hutIGC operon expression (291).
Nitrogen limitation or growth with histidine as the sole nitrogen source induces expression of the hut genes in K. aerogenes and some serovar Typhimurium strains (49, 336), but not in serovar Typhimurium LT-2 (305). This induction in K. aerogenes requires the Ntr response (393), and more specifically Nac (31). Nac activates hutUH expression (290, 460, 462) from the same transcription start site as that used during carbon-limited growth (351). Serovar Typhimurium lacks nac (30); however, Nac induces the serovar Typhimurium hut genes when they are placed in K. aerogenes (35). The binding site for Nac in the K. aerogenes hutUH promoter region is centered at about –64 bp, relative to the transcription start site (163, 385). Nac facilitates the binding of RNA polymerase (163).
Mutations in several regulatory (cis-acting) loci in serovar Typhimurium have been useful tools in analyzing hut expression (49, 305, 483, 485). Alterations in hutP permit utilization of histidine as a nitrogen source (305), or eliminate hutUH expression (49). The hutP locus probably defines the hutUH promoter (485). Mutations in hutR permit hutUH expression with glucose, although expression is still catabolite-repressed (49). These mutations are also probably in the hutUH promoter. In contrast, mutations in the hutQ locus result in constitutive hutUH expression and probably define the hutUH operator (49). The hutM locus defines a region that is necessary, but not sufficient, for utilization of histidine as a carbon source (305). Alterations in this locus, which probably defines the hutIGC promoter, affect synthesis of HutI, HutG, and HutC (177, 483).
Histidine transport has been extensively studied only in serovar Typhimurium, which appears to contain at least five histidine transport systems. The best studied is an ABC-type system consisting of the periplasmic binding protein HisJ (Kd, 0.11 μM) and three membrane components, HisQ, HisM, and HisP (5, 6, 267). It has been a model for the ABC systems. A second periplasmic binding protein for histidine was originally called HisK and subsequently called ArgT (the LAO protein) (histidine Kd, 1.5 μM) (5, 6, 425). ArgT also interacts with the HisQ-HisM-HisP complex (266, 268). HisJ and HisK account for 95% and 5%, respectively, of the histidine-binding activity by proteins released by osmotic shock (5). The HisJ system is the high-affinity system (originally called the HisP system) that was identified as one of two kinetic components in the first study of histidine transport (4). The low-affinity system was called AroP, which has been subsequently well characterized; aromatic amino acids inhibit histidine transport by this system (4, 9). The characteristics and regulation of AroP are discussed in the sections on aromatic amino acid degradation and aromatic amino acid synthesis. An aroP hisP double mutant, which eliminates the HisJ, ArgT, and AroP systems, still has two kinetic components of histidine transport: X, which is inhibited by aromatic amino acids, and Y, which is insensitive to aromatic amino acids (9). Histidine transport has also been studied with membrane vesicles (i.e., systems independent of binding proteins), and these studies also suggest the existence of components both inhibited by, and insensitive to, aromatic amino acids (282). In serovar Typhimurium LT-2 derivatives that can utilize histidine as a nitrogen source, aroP and hisP mutants grow normally, while an aroP hisP double mutant grows twice as slowly (266). The remaining systems, presumably X and Y, appear to be reasonably high-capacity systems. X, Y, and the membrane vesicle systems have not been genetically identified or further characterized.
The genes for the binding protein-dependent systems are clustered. The gene order is argT-hisJQMP. Nitrogen limitation induces synthesis of HisJ (threefold), HisP, ArgT (sixfold) (7, 201, 268) and transcription of argT and hisJQMP (8, 452, 453, 502, 587). The nitrogen control of argT is much greater than that of hisJQMP (8, 268, 452, 502). Ntr control requires NtrC and can be enhanced by constitutively active NtrB (8, 268, 502, 587). A σ54-dependent promoter for argT has been identified and functionally analyzed (8, 453). Functional NtrC binding sites have not been identified (8), and footprinting with phosphorylated NtrC, which should identify these sites, has not been done. There is still some controversy as to whether hisJ is regulated by the Ntr system (8, 201, 452, 502).
It has also been suggested that hisJQMP transcription during nitrogen-limited growth may initiate at the argT promoter (452), which is consistent with the greater protection against S1 nuclease of the complete DNA probe by RNA from nitrogen-limited cells (see figure 5 in reference 8). These regulation studies have been complicated by the use of strains with artificially high hisJ expression (the result of an alteration that allows use of D-histidine as a source of L-histidine; e.g., see reference 8), and the likelihood of additional uncharacterized promoters. These transport systems were first studied in cells that were not nitrogen limited (4, 6), which is sufficient to imply Ntr-independent regulation. Furthermore, argT expression is induced by growth into stationary phase (8) and carbon limitation (453, 502), which implies the existence of additional Ntr-independent regulatory elements.
Proline can be the sole source of carbon, energy, or nitrogen for E. coli, serovar Typhimurium, and K. aerogenes. Proline is also an osmoprotectant in a high-osmolarity medium. However, proline catabolism would not be expected to contribute to proline accumulation, and it does not participate in proline homeostasis with moderate or high osmolarity (93). On the contrary, high osmolarity inhibits proline catabolism (123). Proline catabolism is important for various functions in other organisms, such as the symbiotic association of Rhizobium meliloti (234), the flight of Drosophila melanogaster and other insects (195), p53-mediated apoptosis (384), oxygen radical generation (113), and schizophrenia in humans (see references cited in reference 271). In addition, proline catabolic enzymes in eukaryotes may participate in control of compartmental redox states (378). Proline may be important in these organisms because only a few reactions are required to generate a citric acid cycle intermediate, electrons for electron transport (from FADH 2), and NADPH. The point is that proline catabolism may have other unsuspected functions in bacteria.
Proline is catabolized to glutamate in three steps (Fig. 9). Proline dehydrogenase (also called proline oxidase) oxidizes proline to Δ1-pyrroline-5-carboxylate (P5C) with reduction of FAD. In the second half of the reaction, membrane-associated proline dehydrogenase donates electrons from FADH2 to the quinone pool and subsequently to oxygen or alternative electron acceptors. Proline utilization consumes oxygen more rapidly than the catabolism of any other amino acid (133). The reason for this high capacity is not obvious. P5C is spontaneously converted to glutamic-5-semialdehyde, although an enzyme may enhance this reaction in vivo. P5C dehydrogenase oxidizes glutamic semialdehyde to glutamate with reduction of NAD. Proline catabolism requires the multifunctional PutA, which converts proline to glutamate, and PutP, a proline transporter (108, 312, 405, 406).
PutA catalyzes the proline and P5C dehydrogenase reactions and perhaps also glutamic semialdehyde formation. PutA also has autokinase activity and is a transcriptional regulator that mediates induction by proline. The latter functions are considered in the section on regulation. PutA contains approximately 1,300 residues (3, 168, 180, 279, 311, 444). Sequence analysis suggested that residues 340–590 and 650–1130 contain proline dehydrogenase and P5C dehydrogenase, respectively (279). This has been confirmed for proline dehydrogenase activity, which is within a protein fragment containing the first 669 residues of PutA (548). The third domain (residues 261–612) of this fragment contains all the proline dehydrogenase active-site residues and the FAD-binding site (271).
PutA from E. coli appears to be dimeric (53, 444, 548), while the serovar Typhimurium enzyme exists in a monomer-dimer equilibrium (311). The difference may be a function of the purification method. The dimerization determinants have been localized to residues 1–47, which has been proposed to assume a ribbon-helix-helix structure (172). In homologous proteins, the helix-helix motif participates in dimerization. Chymotrypsin cleaves PutA at residue 58, which is just after the dimerization determinants (172).
FAD is required for proline dehydrogenase activity. FAD is the only redox center for proline dehydrogenase (28), but it is not covalently bound. Purified PutA contains about 0.8 mol of FAD per subunit, and has a Km for FAD of 0.15 μM (52). The FAD-binding site has been determined within the structure of the proline dehydrogenase domain from residues 216–612 (271). Soluble PutA has proline dehydrogenase activity with artificial electron acceptors, which oxidize FADH2. However, in vivo the electrons from FADH2 are donated to the electron transport chain with oxygen as the terminal electron acceptor, as shown by proton translocation activity (1), stoichiometric oxygen consumption and P5C formation (132), absence of hydrogen peroxide generation (309), the requirement for cytochromes (1) and quinones (515), and cyanide inhibition (168, 444, 515). Proline catabolism can also use alternate terminal electron acceptors (295), presumably after transfer of electrons to the quinone pool.
Proline dehydrogenase activity is membrane-associated (1, 132, 168, 309, 311) and it has been proposed that the membrane-associated form is monomeric (unpublished result cited in reference 515) and that the soluble form is dimeric (311). Hydrophobic regions containing residues 160-170, 770–820, and 1205–1220 have been suggested to contribute to membrane binding (279), but this has not been tested. Although PutA1–669 has proline dehydrogenase activity, it does not associate with membranes (548). FAD reduction increases PutA hydrophobicity (363, 515), alters PutA’s structure outside of the FAD active site (52, 586), and is required for membrane association (52, 168, 515, 565). Membrane association requires phospholipids, but not a specific protein or quinones (515). This and other evidence (168) argues against a model that repression involved saturation of specific membrane sites (294). Several lines of evidence suggest that membrane association is required for dissociation of PutA from DNA and release of PutA-dependent repression (discussed below). It has been noted that the catalytic cycle should result in release of PutA from the membrane, and therefore it was proposed that binding to the membrane is irreversible (52). However, regulation is likely to be more complicated, since induction not only requires proline and FAD reduction, but also oxygen or an alternate electron acceptor, and PutA mediates this control (294, 295). The requirement for an electron acceptor has not been explained.
The PutA enzyme is highly specific for L-proline and it has no activity toward D-proline or other amino acids (309, 444). The apparent Km for proline have been reported from 1 to 100 mM. The reported Kms from different laboratories seem to suggest that the Km increases during purification. The lowest reported Km for proline is 1.25 mM and is from membrane-containing cell extracts (132). The apparent Kms for PutA assayed from membranes isolated with different buffers range from 2.7 to 18 mM (1). The reported Kms for purified soluble PutA range from 50 to 100 mM (309, 444, 548). Membrane association lowers the Km by about 40% (309). The midpoint proline concentration required for FAD bleaching or altered proteolytic susceptibility is about 0.1 mM, which suggests that proline binds PutA better than would be indicated by the apparent Km (52). Events after FAD reduction may affect the apparent Km or perhaps PutA’s structure. L-Lactate, D-lactate, and pyruvate inhibit proline dehydrogenase with Kis of 1.4, 2.1, and 3.3 mM, respectively (444).
The conversion of P5C to glutamic semialdehyde can be spontaneous. However, the equilibrium constant for this reaction is unknown (309). The likely P5C dehydrogenase domain, residues 650 to 1130, is homologous to proteins of the aldehyde dehydrogenase superfamily, which implies that glutamic semialdehyde, and not P5C, is the substrate (279). The weak homology of residues 228–358 to serine-pyruvate aminotransferases was used to suggest that PutA catalyzes glutamic semialdehyde production (279). Aspects of the structure of the active site of PutA1–669 also suggested that PutA converts P5C to glutamic semialdehyde (271). The intermediates of proline catabolism appear to be channeled within PutA because proline dehydrogenase catalysis is fourfold faster than that of P5C dehydrogenase with exogenous substrates, and P5C dehydrogenase activity is 14-fold faster with endogenous than with exogenous P5C (514).
The ratio of proline to P5C dehydrogenase is constant during purification, which provided biochemical evidence that one polypeptide contained both activities (311). The Kms for exogenous P5C and NAD are 11 and 0.1 mM, respectively (309). As mentioned above, exogenous P5C is a poor substrate compared with endogenous P5C (514), which may account for the high Km. Little else is known about the NAD-dependent P5C dehydrogenase.
PutA also autophosphorylates on serinyl, threoninyl, and tyrosyl residues (365). Phosphorylation does not affect PutA’s enzymatic activities (365). The function of phosphorylation will be discussed in the section on regulation.
Utilization of proline as a carbon or nitrogen source requires the PutP transport system (Fig. 9) (405, 567). PutP is also the major transport system that supplies proline to a proline auxotroph grown in medium with optimal or medium osmolarity (310). ProP also contributes to transport of proline in an auxotroph, but it only accounts for about 5% of the transport activity (310). ProU can also transport proline in an auxotroph, but only in medium with high osmolarity (94). The primary functions of ProP and ProU are in proline and glycine betaine accumulation in high-osmolarity medium and will not be considered further. A proline auxotroph lacking all three systems can be supplemented with proline, which implies a fourth mechanism of proline transport. A derivative of a putP mutant that can utilize proline as sole nitrogen source apparently altered the LIV-I transport system so that it could transport proline (122). This might represent the fourth proline transport system. A useful review of work on proline transport prior to 1988 has been presented (566).
The PutP system contains a single protein that catalyzes Na+-proline cotransport (58, 78, 503, 566). It was initially considered a model system for the study of H+-proline symport. The Na+ requirement was difficult to show, as evidenced by initial studies that indicated inhibition by Na+ (282). However, this was shown to result from preloading vesicles with Na+, which inhibited Na+ influx (503). Direct evidence for Na+-proline and Li+-proline symport has been provided, but these studies were unable to detect any H+-proline symport (78).
The putA and putP genes are adjacent but divergently transcribed. Figure 10 provides a summary of regulatory sites in the putP-putA intergenic region. There is general agreement that the transcription start site for putA is about 40 bases from the start site of translation in all organisms examined to date (80, 180, 331, 332). There is less agreement concerning the start sites for putP. Three major sites cluster at −130 in E. coli, and one probably corresponds to P2 in Fig. 10 (332). A single major site at −400 has been reported from serovar Typhimurium (180, 364), and sites at −133 and −185 from K. aerogenes (80). However, the scheme represented in Fig. 10 is suggested for all three organisms because of (i) the similar homology of the intergenic region in these organisms (80), (ii) the detection of these transcripts in all three organisms (80), and (iii) the location of the proposed or verified regulatory sites in relation to the start sites (described below). There is a relatively high basal level of expression, which is not surprising considering PutA’s function as a repressor. The uninduced and induced levels of PutA have been calculated to be about 1,000 and 10,000 molecules per cell, respectively (294).
Proline, and not a metabolite derived from proline, induces expression of both genes in the three enteric organisms considered here (108, 123, 133, 295, 405). PutA is a transcriptional repressor that mediates proline-dependent induction of expression. The DNA-binding determinants are located within the first 47 residues of PutA, as demonstrated by the DNA-binding of PutA1–47 in vivo and in vitro (172). This small protein fragment binds DNA cooperatively, with a Kd of 15 nM, which is about three times lower than the Kd for wild-type PutA (172). Mobility shift assays indicate that purified PutA binds several sites within the putP-putA intergenic region (364). Five sites have been identified based on a repeated 27-bp sequence with dyad symmetry (labeled O1 to O5 in Fig. 10) (364). This has not been verified by either footprinting or genetic alteration. Repression of putA expression requires sequences that are at least 180 bases upstream from the translational start site (331), which may implicate O3, O4, or O5 in such repression.
PutA is not sufficient to repress putA transcription with purified components (unpublished result stated in reference 356). PutA-dependent repression requires integration host factor (IHF) in vivo (356). Footprinting experiments have identified the IHF sites (356). IHF-A’s location would seem to suggest that it is more important than IHF-P (Fig. 10). In addition to IHF sites, two sites for DNA bends have been identified (356). These results suggest the formation of a large nucleoprotein complex.
Alterations that prevent induction by proline invariably alter proline dehydrogenase activity, which suggests that dehydrogenase activity is required for derepression (322). This is consistent with the observation that constitutive alterations span the entire putA gene, that is, they can be outside the DNA-binding region (312). Proline is insufficient for derepression in vitro (52, 363). Instead, proline and either an artificial electron acceptor or membranes are required for derepression (363). The redox state of FAD also does not affect binding to DNA (28), and binding to DNA and membranes is mutually exclusive (323). This has led to the hypothesis that membrane association is required for derepression (28).
In addition to proline, oxygen or alternate electron acceptors induce putA expression in serovar Typhimurium (295). However, E. coli cannot utilize proline as carbon source for anaerobic growth with fumarate or nitrate as electron acceptors, which suggests that alternate electron acceptors do not induce expression (G. Baker and J. M. Wood, unpublished results cited in reference 566).
PutA has autokinase activity, and phosphorylation inhibits binding to DNA (365). Proline stimulates dephosphorylation of soluble PutA but has no effect on membrane-bound PutA (365). The former effect probably does not occur in vivo, since proline and FAD reduction will promote membrane association. Perhaps phosphorylation prevents PutA-dependent repression if PutA is released from the membrane, although an alternate explanation has been proposed (365).
A novel regulatory mechanism was suggested when a mutant lacking the leucyl,phenylalanyl-tRNA-protein transferase was shown to have fourfold more proline dehydrogenase (110). The transferase modifies proteins and targets them for degradation (531). However, the transferase does not modify PutA (445), but it was shown that the parental strain of the transferase mutant has a mutation near or in putA that results in premature cessation of induction, whereas the transferase mutant does not have this mutation (109). Therefore, this seemingly novel regulatory mechanism only further confirmed PutA’s regulatory function.
Glucose represses putA and putP, and cyclic AMP and CRP can alleviate this repression (108, 179, 295, 405). Mutations in cya or crp lower putA and putP expression (405). Putative CRP-binding sites have been deduced from sequence analysis (80, 180, 331, 364), but binding of CRP to these sites has not been verified biochemically or genetically. The only information available about these sites is that a deletion analysis located the CRP sites for putA and putP expression within 107 and 234 bases of their respective start codons (331). This implicates the proposed CRP-A and CRP1 sites, respectively, in catabolite repression (Fig. 10).
Nitrogen limitation in proline-containing medium induces proline dehydrogenase synthesis fourfold in K. aerogenes (394). Nitrogen limitation increases expression from the P1 promoter in K. aerogenes, which is the same promoter as that used during carbon limitation (79, 80). Nac is required for growth with proline as the nitrogen source (31) and induces proline catabolic genes in K. aerogenes (324). The Nac-binding site has been identified and is appropriately located to activate the P1 promoter of putP, but not the P2 promoter or the putA promoter (79). Nitrogen limitation does not induce either PutA or PutP synthesis in E. coli (394), although overexpression of NtrC, which should induce Nac, increases putP-lacZ expression twofold, but decreases putA-lacZ expression (331). Nitrogen limitation modestly represses proline transport (presumably via PutP) in serovar Typhimurium (142). Serovar Typhimurium lacks both Nac and binding sites for Nac (79). Nonetheless, serovar Typhimurium can still utilize proline as the sole nitrogen source, presumably because of high basal putA expression.
These amino acids are grouped together because serine is a product of glycine catabolism, which in turn is a product of threonine catabolism. The pathways of their catabolism are summarized in Fig. 11. E. coli can utilize L-serine as a carbon or nitrogen source, but optimal growth requires supplemental amino acids. Serovar Typhimurium and K. aerogenes can apparently use L-serine as a carbon or energy source without supplementation. Amino acid supplementation improves growth, at least for K. aerogenes. These bacteria can use glycine or threonine as a nitrogen source, although efficient threonine utilization requires amino acid supplementation.
Serine is toxic in minimal medium, and supplemental amino acids prevent this toxicity (; reference 184 and references therein). Serine inhibits threonine deaminase (404, 539), homoserine dehydrogenase I (184), and presumably cystathionine β-lyase (MetC) (51). Serine also inhibits glutamine synthetase (277). Note that cysteine inhibits many of the same enzymes as serine. Glycine, methionine, leucine, and high temperature exacerbate the toxicity (96, 543). In addition to inhibition of several enzymes, it has been proposed that high pyruvate concentration resulting from enhanced serine catabolism restricts isoleucine synthesis by an acetohydroxy acid synthase isozyme (303). Serine toxicity may be a combination of inhibition of specific enzymes compounded by the effect of high pyruvate. However, pyruvate by itself is not toxic. In addition, environmental factors can affect serine toxicity, and serine-dependent inhibition of homoserine dehydrogenase I requires physiological K+ and pH (184). Isoleucine overcomes serine toxicity in nitrogen-rich (i.e., ammonia-containing) minimal medium, whereas methionine, but not isoleucine, overcomes toxicity when serine is used as the sole nitrogen source (457).
Several different enzymes have the potential to degrade serine to pyruvate and ammonia. E. coli contains three serine deaminases (dehydratases): SdaA (LSD1) (L-serine Km, 2.7 mM; D-serine Ki, 1.4 mM; L-cysteine Ki, 0.9 mM), SdaB (LSD2), and TdcG (82, 304, 440). Cystathionine β-lyase (MetC), tryptophanase (TnaA) (Km, 160 mM), and the B subunit of tryptophan synthase also have serine deaminase activity (51, 92, 348, 486). Serine can be degraded as a carbon source in aerobic minimal medium if glycine and leucine are also present (346). This is consistent with early observations of aerobic control of serine deaminase synthesis that showed induction by a casein hydrolysate, and more specifically by glycine and leucine (219, 369). Growth with serine (and glycine and leucine) as a carbon source is eliminated through insertional inactivation of sdaA (511), impaired in mutants with low SdaA (269, 344), and enhanced with elevated SdaA (345, 511, 534). Growth with serine as a carbon source is also abolished by certain lesions in transport systems (discussed below). In addition, high levels of SdaB (510) and MetC (51) also stimulate growth, which indicates their potential for serine catabolism.
Growth with serine as a nitrogen source is diagnostic of a functional Ntr response. Such growth requires supplemental methionine but not isoleucine. The enzymes that degrade serine as a nitrogen source are not known. A microarray analysis of Ntr-regulated genes indicates that Ntr regulators do not control any possible serine catabolic or transport protein (587). This is consistent with the observation that E. coli grown in ammonia-containing (i.e., nitrogen-rich) medium adapts immediately to growth with serine as a nitrogen source, which suggests the presence of serine catabolic enzymes for cells grown in nitrogen-rich medium (346). Serine as a nitrogen source does not activate Ntr genes as effectively as other nitrogen sources. Serine induces glnA, but only weakly activates other Ntr genes (410). Even though Ntr regulators do not control serine catabolism, they do control serine synthesis. Nitrogen limitation induces Nac, which represses the first enzyme of serine synthesis (40). This regulation may integrate the synthesis of components required for nucleotide synthesis (i.e., C1 units and glycine) with nitrogen limitation.
SdaA is the major serine deaminase during aerobic growth in minimal medium (511). Leucine or glycine, but not serine, increases SdaA activity or sdaA transcription in E. coli (219, 369, 511). Lrp represses sdaA expression, and leucine partially reverses this repression (534). Other factors also affect sdaA expression or SdaA activity, and their modes of action are poorly understood and possibly indirect. Nitrogen limitation increases E. coli SdaA activity 70% (219). One mutation that increases sdaA expression was a lesion in what was originally called wyb, subsequently called ssb, and finally termed cpx (although evidence for the latter identity has not been presented) (219, 304, 320, 343, 345, 525). This lesion affects a variety of growth properties. Lesions in torA, nuoM, dms, and glpC impair growth with serine (and glycine and leucine) as the carbon source, although SdaA synthesis and activity are not affected in glucose-containing medium, and strains with these lesions are not stable (269). The products of these genes are involved in anaerobic redox functions. These observations may be related to the observation that anaerobic growth in alkaline medium induces sdaA (583). High temperature and heat shock also induce serine deaminase activity, and transcription appears to be affected (303, 339). This induction does not require σ32 (303). Not surprisingly, ethanol also induces serine deaminase activity (27). Glucose, however, does not repress serine deaminase activity (which is undoubtedly mostly SdaA) and medium with glucose has the highest activity (219). These results imply that catabolite repression does not control serine deaminase synthesis. Nothing is known about glycine-dependent induction of sdaA expression; however, the only known regulatory effect of glycine is on regulators of the glycine cleavage enzyme (described below).
The serine deaminases are notoriously unstable, which has been known since their first assay in 1938 (145, 369). Iron and a reducing agent are required for full activity of SdaA and SdaB (347, 510). SdaA is the best characterized serine deaminase and is probably prototypical considering there is greater than 75% amino acid identity between SdaA, SdaB, and TdcG. SdaA has an unstable [4Fe-4S] cluster, which is required for activity (82). Protein cysteines bind three irons, and substrate serine binds the fourth (82). Solvent exposure of one iron undoubtedly contributes to Fe-S cluster instability and enzyme lability. Various environmental factors affect SdaA activity, and reversible Fe-S assembly may mediate these effects. DNA-damaging agents (acting through the SOS response) and other stresses increase SdaA activity (339), but do not activate sdaA expression, which suggests that they affect the enzyme activation mechanism (511). Some anaerobically induced redox functions may be necessary for Fe-S cluster assembly (269).
SdaB accounts for 50% of serine deaminase activity in cells grown in nutrient broth (510). The sdaB gene is part of the sdaCB operon, and the first gene specifies a serine transport system (472). The sdaB mRNA is poorly translated (473). Loss of CRP reduces expression of the operon and SdaB activity 20- to 30-fold, although glucose reduces expression only twofold (472, 473). Leucine induces sdaCB expression up to threefold, and Lrp mediates this control (472, 473). sdaB expression and SdaB activity are not affected by DNA-damaging agents, high temperature, or anaerobic growth (473, 510).
The anaerobically induced serine deaminase, TdcG, has recently been purified and characterized as a His-tagged derivative (54a). It is homodimeric and appears to have two oxygen-labile Fe4-S4 clusters, which are necessary for activity. It has an apparent Km for L-serine of 4.8 mM, but does not react with threonine. TdcG is specified by the last gene of the tdcABCDEFG operon. The tdc gene products degrade serine and threonine anaerobically. Threonine and low glucose induce expression, with the latter requiring cyclic AMP and CRP. This operon and its products will be discussed more thoroughly in the section on threonine catabolism.
Regulation of assayable serine deaminase in K. aerogenes is similar, but not identical, to that in E. coli (547). Leucine, glycine, and threonine induce activity, whereas nitrogen limitation and exogenous serine do not. K. aerogenes has at least two serine-degrading enzymes: a serine deaminase (presumably SdaA) and a nonspecific isoleucine-sensitive threonine dehydratase. Elevated synthesis of either enzyme stimulates growth with serine as carbon or nitrogen source.
E. colicontains two major serine transport systems and apparently at least one minor system. The major system in minimal medium is a Na+-dependent serine/threonine cotransport system (186). Threonine-inhibitable serine transport is readily detectable with membrane vesicles isolated from cells grown in minimal medium (282), and indeed with whole cells grown in minimal medium (526). A mutant lacking this system is resistant to serine toxicity in minimal medium and fails to utilize serine as a carbon source (357). The gene for this transport system is sstT (sodium-serine-threonine transport), the former ygjU (358). Purified SstT in liposomes catalyzes threonine-inhibitable Na+-dependent serine transport (254). Initial studies indicated that SstT synthesis is constitutive (527); however, growth in nutrient broth represses synthesis, and tryptophan appears to be the only repressing amino acid (186, 358). A nhaB mutant, which lacks a Na+/H+ antiporter, also fails to utilize serine as a carbon source, presumably because of the establishment of an inadequate Na+ gradient, which is required for serine transport (220). The growth of this mutant is defective in alkaline pH (220), which is consistent with a proposed function of serine catabolism in cytoplasmic acidification (see below). The defect in an nhaB mutant does not appear to be consistent with results from other studies, which suggest that NhaB functions below pH 8.0 (435). However, the growth media differ substantially with respect to sodium content. It is possible that the results are correct for the medium used, or that strain differences affect at least one of the three Na+/H+ antiporters.
Growth in nutrient broth appears to repress the Na+-dependent serine system and to induce a threonine-insensitive H+-serine cotransport system (185, 186). Leucine or methionine is the inducing component in broth (185, 186). The sdaC gene almost certainly codes for this transport protein. Expression of sdaC parallels that of the H+-serine system, and both are induced in nutrient broth and modestly by leucine in minimal medium (186, 472). Furthermore, sdaC on a plasmid elevates threonine-insensitive serine transport (472). Its expression depends almost completely on CRP (472); however, glucose only modestly affects expression (185, 472). Mutationally elevated, leucine-inducible, H+-dependent serine transport can suppress the defect in an nhaB mutant (249). The induction by leucine suggests that the mutant has enhanced sdaC expression. SdaC is homologous to TdcC (472), which is an anaerobically induced H+-serine/threonine cotransporter (described in the section on threonine catabolism). Multicopy tdcC can suppress loss of sstT and permit growth with serine as a carbon source (357). Serine inhibits L-alanine transport by the major transport system for L-alanine, which is the LIV-I system for branched-chain amino acids (419). However, L-alanine did not inhibit serine uptake, perhaps because rapid metabolism made serine transport difficult to assay (419). Nonetheless, serine inhibited leucine uptake, which is the preferred substrate for the LIV-I system and competed with leucine for binding to the leucine-binding protein (403). The only evidence that the LIV-I system transports serine is competition assays, and it is possible that serine bound to the binding protein is not actually transported.
The unusual induction requirements for the serine catabolic and transport proteins do not readily suggest catabolic functions for the serine deaminases. Inducing conditions are probably most often encountered in complex medium. Perhaps the serine deaminases prevent serine toxicity in such situations. This is consistent with the observation that serine is the first amino acid degraded in an amino acid mixture (401). It has also been proposed that serine catabolism acidifies the cytoplasm of anaerobic cells grown in alkaline medium (583). During anaerobic growth, serine will be nonoxidatively degraded to ammonia and pyruvate. The basic ammonia will be released, and subsequent pyruvate degradation will generate acetic and formic acids. This is consistent with the alkaline induction of sdaA, and the requirement of the SstT transport system for Na+/H+ antiport, which is essential for growth in alkaline medium.
There have been no developments on D-serine catabolism since this topic was reviewed by McFall and Newman (304). D-Serine is toxic to cells grown in minimal medium and it inhibits synthesis of L-serine and pantothenate (and perhaps some enzymes inhibited by L-serine). D-Serine dehydratase, the product of dsdA, degrades D-serine to pyruvate and ammonia. The dehydratase is monomeric (47,000 Da), requires pyridoxal-5'-phosphate, and has a Km for D-serine of 0.32 mM. Activation requires the LysR-like transcriptional regulator DsdC complexed to D-serine, and this activation is enhanced by, but does not require, cyclic AMP and CRP. The dsdXA operon is divergently transcribed from the dsdC operon; DsdX is a putative membrane protein with no known function. The dehydratase appears to prevent D-serine toxicity and degrade D-serine as a carbon or nitrogen source. L-Serine is more rapidly racemized than most amino acids, which may account for the presence of D-serine dehydratase.
The use of glycine as a sole nitrogen source has been studied with E. coli. Leucine stimulates such growth, and the stimulation could not be entirely attributed to leucine deamination (338). Labeling studies indicate that glycine is converted to serine, which is degraded to pyruvate (340). Mutants lacking either the glycine cleavage enzyme (GCV) or serine hydroxymethyltransferase (GlyA) cannot use glycine as a nitrogen source (340). GCV cleaves glycine to NH3, CO2, and C1 (as N5,N10-methylenetetrahydrofolate). The C1 unit is required for serine formation. GCV is a four-protein complex from bacteria to mammals (250, 498). In E. coli, the gcvTHP operon specifies three proteins, while lpd specifies a fourth component that is also part of the pyruvate and α-ketoglutarate dehydrogenase complexes (498, 501). GlyA forms serine from N5,N10-methylenetetrahydrofolate and glycine. GlyA contains pyridoxal-5'-phosphate and zinc (247). Both GlyA and GCV can be considered biosynthetic enzymes. GlyA synthesizes glycine, while GCV contributes to the formation of C1 units. The control of these enzymes is thoroughly reviewed in Chapter Regulation of Serine, Glycine, and One-Carbon Biosynthesis. The CycA protein (also called the glycine-alanine system) transports glycine by using the proton motive force as the energy source. Its properties and regulation are discussed in Chapters Regulation of Serine, Glycine, and One-Carbon Biosynthesis and Biosynthesis of Glutamate, Aspartate, Asparagine, l-Alanine, and d-AlanineBiosynthesis of Glutamate, Aspartate, Asparagine, l-Alanine, and d-Alanine (in the section on alanine transport). Catabolic aspects are briefly summarized here.
Growth with glycine as the nitrogen source is slower than with ammonia (340). Furthermore, glycine inhibits glutamine synthetase (277) and, therefore, glycine as a nitrogen source should induce the Ntr response. Nitrogen limitation and Ntr regulators do not affect gcvTHP or glyA expression but modestly induce cycA expression, which encodes the major glycine transport protein (587). Instead of Ntr control, glycine regulates expression of the glycine catabolic enzymes. Glycine controls gcvTHP expression through GcvR and GcvA. Without glycine, the GcvR-GcvA complex represses expression. Glycine binds GcvR, disrupts the interaction between GcvR and GcvA, and GcvA, which is homologous to LysR, activates gcvTHP expression (196). In addition to these regulators, a PurR-hypoxanthine/guanine complex represses gcvTHP (196). Glycine may indirectly stimulate PurR-dependent repression, since glycine is a substrate for purine synthesis, which consequently may increase hypoxanthine and guanine synthesis. Lrp is essential for gcvTHP expression, but its role appears to be structural (499). Leucine does not affect the Lrp-dependent activation (342). Glycine as the sole nitrogen source moderately represses glyA (340) and it is activated by MetR-homocysteine and repressed by PurR-hypoxanthine/guanine (283). Glycine has the potential indirectly to control both regulators. Glycine may stimulate purine synthesis and PurR-dependent repression. Glycine degradation by GCV will produce C1 units, which may deplete homocysteine and prevent MetR-dependent activation of glyA. In addition to these factors, Lrp modestly represses glyA (342).
Four different enzymes initiate pathways of threonine utilization: biosynthetic threonine deaminase (IlvA), threonine aldolase (LtaE), threonine dehydrogenase (Tdh), and biodegradative threonine dehydratase/deaminase (TdcB). IlvA is the first enzyme of isoleucine synthesis and is one target of serine and cysteine toxicity. A serovar Typhimurium mutant in which IlvA is insensitive to isoleucine inhibition utilizes threonine as a nitrogen source (56). E. coli mutants with a similar phenotype have not been described.
Threonine Aldolase.
Threonine aldolase (LtaE) cleaves threonine (Km, 2.85 mM) and L-allo-threonine (Km, 0.22 mM) to acetaldehyde and glycine (281). An E. coli ltaE glyA mutant grows four times more slowly than a glyA mutant in minimal medium, which indicates that threonine aldolase synthesizes glycine in vivo (281). However, GlyA is 70 times more active than LtaE in a crude extract (281). Purified GlyA also has extremely weak threonine aldolase activity (449). The substrates threonine and L-allo-threonine as sole carbon sources in minimal medium or as supplements in nutrient broth do not affect LtaE synthesis, which argues against a catabolic function (281).
The Threonine Dehydrogenase Pathway.
Threonine dehydrogenase (Tdh) catalyzes the NAD-dependent oxidation of threonine to 2-amino-3-ketobutyrate, which 2-amino-3-ketobutyrate CoA ligase (Kbl) then cleaves to acetyl-CoA and glycine. Tdh is a tetramer with 35-kDa subunits (48). The Km for threonine is 1.4 mM, and the enzyme has a pH optimum of 10.3 (48). An unidentified metal and thiols activate Tdh activity (91). Tdh is homologous to known zinc-dependent enzymes (19). Kbl is a dimer of 42-kDa subunits (321), it contains pyridoxal-5'-phosphate, and the Kms for acetyl-CoA and glycine are 59 μM and 12 mM, respectively (321). (The reaction is assayed in the reverse direction.). This pathway is reversible but appears to function only in threonine catabolism in vivo (296). The E. coli genes encoding Tdh and Kbl form the kbl-tdh operon. Leucine, but not threonine, induces kbl-tdh expression (341, 417), while Lrp represses the kbl-tdh operon 20-fold. Leucine antagonizes this repression, and results in a 10-fold induction (417). The promoter and Lrp-binding site have been determined (18, 417).
The Tdh-Kbl pathway serves three functions: a source of glycine; threonine utilization as a nitrogen source; and threonine utilization as a carbon source. The pathway was first discovered as a threonine-dependent pathway of glycine synthesis (136, 341). Leucine, but not threonine, induces Tdh activity (341). An insertion in tdh prevents conversion of threonine to serine (in a strain blocked in serine synthesis from 3-phosphoglycerate) (407). Optimal expression of kbl-tdh requires a combination of leucine, arginine, lysine, threonine, and methionine (407). E. coli can slowly degrade threonine as a nitrogen source but only if leucine is also present (390). Labeling studies and the inability of glyA and gcvTHP mutants to utilize threonine as a nitrogen source indicate that glycine and serine are obligatory intermediates in threonine degradation (390). With threonine as the nitrogen source, glycine and serine are almost entirely derived from threonine, not from glucose (390). Utilization of threonine as a nitrogen source correlates with Tdh, not TdcB, activity (390). A mutant with very high Tdh activity does not require leucine for such growth (390). Ntr regulators do not directly affect expression of the kbl-tdh operon during nitrogen-limited growth (587). Nonetheless, the slow growth with threonine as a nitrogen source probably requires optimal glutamine synthetase activity and therefore induction of the Ntr response. E. coli can use threonine as a carbon source. During such growth, the acetyl-CoA generated is necessary for growth but the glycine is not (76). Several subsequent reports indicate that threonine utilization requires high levels of Tdh and Kbl, obtained through mutation or overexpression of the kbl-tdh operon on a multicopy plasmid (17, 47, 408). These mutants are hypersensitive to valine, presumably because of threonine and isoleucine depletion (417). Tdh in such strains is no longer leucine inducible presumably because of the loss of Lrp regulation (17, 47).
The Anaerobic Threonine Dehydratase Pathway.
Threonine dehydratase/deaminase (TdcB), also called the biodegradative threonine deaminase, catalyzes the first reaction in a pathway that anaerobically degrades threonine (199, 440). TdcB degrades threonine to α-ketobutyrate and ammonia. α-Ketobutyrate is cleaved to propionyl-CoA and formate, and propionyl-CoA is then converted to propionyl-phosphate, which can drive substrate-level phosphorylation, ultimately producing propionate. An energy-generating function is consistent with the requirement for cyclic AMP-CRP, the tight catabolite repression, and the substrate-level phosphorylation. However, a strain with an insertion in tdcB has no obvious growth defect in a variety of media (164). The genes coding for enzymes that catalyze the second and fourth reactions of this pathway are part of an operon containing tdcB (199). Deletion of these genes results in no discernible phenotype (199).
Threonine dehydratase has been much more intensively studied than other enzymes of this pathway. The induced enzyme appears to be about 2% of soluble protein (480). Purified threonine dehydratase from E. coli and serovar Typhimurium is a homotetramer of 35-kDa subunits (37, 433). The purified E. coli enzyme contains four molecules of pyridoxal-5'-phosphate per tetramer (480), but only two are required for activity (373). The serovar Typhimurium enzyme contains only two pyridoxal-5'-phosphates per tetramer (37). A characteristic property of this enzyme is that AMP is an allosteric activator and is essentially required for activity. The Km for threonine is 4 to 11 mM with AMP, and about 100 mM without AMP (480, 559). Threonine dehydratase also cleaves serine (Km, 5 mM with AMP; 41 mM without AMP) (480). The kinetics are nonsigmoidal (559), but intracellular threonine and serine should not be depleted because of the high Kms. One function of AMP is to counteract inhibition by α-ketobutyrate, a competitive inhibitor (479). The Kactivation for AMP is 2 μM without α-ketobutyrate and 24 μM with α-ketobutyrate (479). AMP shifts the pH optimum from 9.5 to a broad optimum of 7.5 to 10 (204). AMP also affects the state of oligomerization (204) with α-ketobutyrate promoting dissociation, which AMP reverses (479). Synthesis of threonine dehydratase is also subject to catabolite inactivation. Glucose, pyruvate, and other metabolites inactivate threonine dehydratase in whole cells (128). Pyruvate, but not glucose, also inactivates the purified protein (128). The inactivation is irreversible, results in disassociation of the tetramer, and involves covalent attachment of pyruvate (128). This inactivation requires long incubations (several hours) and is distinct from kinetic inhibition of activity during the course of a short enzyme assay. Pyruvate also inhibits threonine dehydratase and binds to sites distinct from those for threonine or AMP (372). Glyoxylate also inhibits and inactivates threonine dehydratase and binds covalently. The effects of glyoxylate and pyruvate are additive and occur by different mechanisms (373, 374, 376). An unusual property of threonine dehydratase is that amino acid composition indicates 1 to 2 mol of tryptophan per subunit, but the DNA sequence indicates no tryptophan codons (99). Purified threonine dehydratase covalently binds exogenous tryptophan (99). These properties distinguish threonine dehydratase from the biosynthetic threonine deaminase (IlvA), which catalyzes the same reaction. Another distinguishing property is that isoleucine inhibits threonine deaminase, but not threonine dehydratase (480, 540). Induced TdcB is 20 to 50 times more active than IlvA (540). The second enzyme of the pathway is 2-keto acid formate-lyase (TdcE), which catalyzes the CoA-dependent cleavage of α-ketobutyrate to propionyl-CoA and formate (199, 442). TdcE can functionally replace pyruvate formate-lyase (199, 442). TdcE and pyruvate formate-lyase are both glycyl radical enzymes that are activated by the same activating enzyme (199). Phosphotransacetylase (Pta) catalyzes the third reaction of the pathway, conversion of propionyl-CoA to propionyl phosphate (199). A propionate kinase (TdcD), which also has acetate kinase activity, catalyzes the last reaction (199).
The genes of the tdcABCDEFG operon specify a transcriptional regulator (TdcA), threonine dehydratase (TcdB), a threonine-serine transporter (TdcC), propionate kinase (TdcD), 2-keto acid formate-lyase (TdcE), and a serine deaminase (TdcG) (54a, 199, 440). The function of TdcF is not known. The tdcR gene, which specifies a regulator of the tdc operon, is adjacent and divergently transcribed. There is a single operon in most strains of E. coli, but in some derivatives of W3110, the tdc operon is not intact. Instead, tdcR-tdcABCD' has been amplified four times, and in a fifth repeat an insertion sequence has replaced part of tdcA, all of tdcBC, and most of tdcD (200, 259). A single transcript and promoter for the complete operon has been detected (166, 199, 442). Glucose represses expression, which requires anaerobic growth in rich medium (121, 464, 540). The most important component of the medium appears to be threonine (121), but other amino acids have also been implicated (121, 206, 585). The pattern of expression has been called multivalent induction (585). Four DNA-binding regulators directly control the tdc operon: cyclic AMP-CRP, IHF, TdcA, and TdcR. Cyclic AMP and CRP mediate the glucose effect, and loss of a CRP site reduces expression (166, 206, 379, 478, 574). IHF is also required for expression, and its binding site is centered at about −100 (574, 575). IHF and CRP bind independently, and both bend DNA (574). The first gene of the operon specifies TdcA, a LysR-like activator. Multicopy tdcA can suppress defects in a strain with an undefined lesion (146), and in several tdcR mutants (178). Indirect evidence suggests that TdcA binds at about −175, but direct evidence for DNA binding has not been obtained (146, 178). TdcR is a 72-residue protein that is also required for tdc expression (178, 465). tdcR expression does not require IHF and does not respond to oxygen or glucose (441, 575). The major mechanism of activation appears to involve realigning CRP and RNA polymerase. The CRP site is centered at −43.5, which is not optimal for an interaction with RNA polymerase (441). Optimal spacing eliminates glucose repression but not the requirement for CRP, the other regulators, or anaerobiosis (441). Therefore, it was proposed that TdcA and TdcR mediate the anaerobic signal (441). Several observations suggest that the topological state of the DNA is important for expression (441, 513, 573). As noted previously, IHF and CRP bend the tdc promoter region (574), and CRP can activate transcription in vitro on a supercoiled template but not a linear template (441). This seems to be consistent with the increase in DNA supercoiling that enhances anaerobically expressed genes (e.g., reference 577). However, factors that relax DNA supercoiling increase tdc expression in vivo (573). The effect of DNA topology on tdc expression is not clear, but it appears that expression requires a properly structured DNA-protein complex.
A variety of factors indirectly affect threonine dehydratase synthesis or tdc operon expression. Alternate electron acceptors (e.g., fumarate) can increase expression (121, 206, 313), but their effects appear to be indirect (77). Mutations in fnr, arcA, pgi and other genes required for carbon flow diminish tdc expression, and it was proposed that a regulatory metabolite does not accumulate in these mutants (77, 313). Perhaps the various amino acids in the growth medium that affect expression (121, 585) contribute to this metabolic control. Cyclic AMP overcomes many of these effects, which is consistent with indirect metabolic control (77). Finally, it has been noted that the tdc operon is the least sensitive of about 1,400 E. coli genes to rifampicin for aerobic cells grown in nutrient broth (468).
Transport.
The three known threonine transport systems, SstT, LIV-I, and TdcC, also transport serine. The first two were described in the section on serine catabolism. In brief, SstT is a Na+-dependent serine/threonine cotransporter, which growth in nutrient broth suppresses. The LIV-I system transports branched-chain amino acids and threonine (419). Leucine (via Lrp) represses this system in minimal medium (342). LIV-I is twice as active as SstT at transporting threonine, at least with the threonine concentration used in the assay (526). The third transport protein is TdcC, specified by the third gene of the tdc operon (512). TdcC is a H+-dependent threonine/serine cotransporter (357, 512). It is regulated, as expected, by anaerobic growth in complex medium (512). Elevated tdcC expression suppresses loss of SstC (357).
Grant MCB0323931 from the National Science Foundation supported the research of L.R. during preparation of this review.
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Module3.4.7
