Amino Acids as Carbon Sources
Chapter
22
ELIZABETH McFALL and ELAINE B. NEWMAN
Most of the 20 "natural" l-amino acids and several d-amino acids are at least potential sources of carbon and energy for members of the Enterobacteriaceae. "Wild-type" laboratory strains are, however, heterogeneous among the various genera with regard to their ability to utilize single amino acids. In some cases genetic information is absent or defective; in some cases it is poorly expressed.
In this chapter we will consider each of the amino acids known to serve as carbon and energy sources for the Enterobacteriaceae, describing the pathways, enzymatic reactions, uptake systems, genetic organization, and regulatory patterns. We will also consider how the cell provides for the multiple functions of particular amino acids in regulating the expression of the corresponding degradative and uptake systems, and we will evaluate the roles of these systems in the cell’s economy. We will be concerned primarily with the l-amino acids. These are what the Enterobacteriaceae see for the most part in their natural environments, and with what most research on amino acid utilization has been concerned. Little is known about enzymatic isomerizations except for those affecting alanine (280), but significant amounts of certain d-amino acids, particularly d-serine, are produced in nonenzymatic isomerizations (86). The catabolism of d-isomers which may serve as utilizable carbon sources for the Enterobacteriaceae is discussed in sections on d-serine and on alanine and d-amino acid dehydrogenase. We have not surveyed degradative enzyme systems which give rise to nonutilizable amines such as cadaverine (254). The function of these pathways is most probably related to maintaining an acceptable intracellular environment, not to carbon and energy metabolism.
For purposes of continuity, the individual amino acids are grouped according to major functional and physiological features of the degradative pathways. We also include a brief discussion of the utilization of mixtures. Some gene locations have been slightly revised from the literature, according to current Escherichia coli (9) and Salmonella typhimurium (241) linkage maps.
With the exception of one of the two l-arginine pathways, degradative pathways for l-arginine, l-glutamine, l-histidine, and l-proline give rise to l-glutamate. This would seem an excellent arrangement. Endogenous l-glutamate itself is readily utilizable for carbon and energy generation via α-ketoglutarate, it maintains much of the carbon skeleton and the C-N bond intact, and it is the focal point of nitrogen assimilation (269). It may serve as a checkpoint, allowing amino acids of five or more carbons to be channeled into anabolic processes if appropriate, before further breakdown occurs. γ-Aminobutyrate, a four-carbon compound, is considered with this group under a separate heading as it is an intermediate in alternate pathways of l-arginine and l-glutamate degradation.
L-Glutamate.
L-Glutamate does not serve as sole carbon source for wild-type E. coli because of inadequate transport (107, 166, 167). L-Glutamate serves as sole nitrogen source but not as sole carbon source for S. typhimurium (153) and as sole nitrogen source for Klebsiella aerogenes (269). Most studies on L-glutamate catabolism have concerned E. coli.
l-Glutamate is of primary importance to the cellular economy in the Enterobacteriaceae because of its central role in nitrogen assimilation and as amino donor in the biosynthesis of other nitrogenous compounds (269). It also appears to be the cell’s first line of defense against osmotic stress. S. Kustu (personal communication) and Csonka (53) have shown that under conditions of osmotic stress in S. typhimurium, synthesis of l-glutamate is enhanced. Kustu (personal communication) found that this increase is essential at moderately elevated osmolarities since, if it is blocked by mutation, the cells do not grow.
There are at least three discrete pathways that can degrade glutamate in the Enterobacteriaceae. From their control patterns, they probably serve separate purposes.
(i) The glutamate dehydrogenase reaction (l-glutamate:NAD+ oxidoreductase, deaminating), NH3 + α-ketoglutarate → l-glutamate, is reversible, but its function seems to be primarily biosynthetic. Its level is high in growth on glucose and low in growth on l-glutamate (109). A gdh mutant still used l-glutamate as carbon source (276).
(ii) The glutamate decarboxylase of E. coli, a pyridoxal phosphate enzyme which converts l-glutamate to γ-aminobutyrate, is formed at high levels at low pH. Wild-type E. coli cannot utilize γ-aminobutyrate as sole carbon source (66), but mutant strains have been isolated that do (see below), and there is no correlation between decarboxylase level and utilization of glutamate (168). Smith et al. (255) suggested that the enzyme may have a role in maintaining physiological pH under acidic conditions, citing Gale’s previous observations on this point. gad loci have been mapped to min 31–34 and min 85 (255) and also to min 82 (155). S. typhimurium does not seem to have a gad gene, and the distribution of the enzyme among the other Enterobacteriaceae appears quite limited (255).
(iii) The primary pathway for use of l-glutamate as carbon source in E. coli was shown by Halpern and his colleagues (109, 170) to be transamination:
l-Glutamate + oxalacetate → l-aspartate + α-ketoglutarate: aspartate aminotransferase
l-aspartate → fumarate + NH4 +: aspartate ammonia-lyase
The oxalacetate needed to continue the reaction is regenerated through the tricarboxylic acid cycle. The lyase (aspartase) is described below (see l-Aspartate). Mutants unable to form the aminotransferase (aspC, min 21) or the lyase were unable to use l-glutamate as carbon source (169). Woods and Guest (301) showed that basal aspartase was similarly repressed in growth with glucose, and that the glucose effect was most probably mediated by cyclic AMP (cAMP)–cAMP-binding protein (CAP). These results clearly demonstrate a catabolic role for the transamination pathway. The same pathway has been shown to exist in K. aerogenes (269).
Although E. coli possesses three separate systems capable of l-glutamate transport, the wild-type levels together are not sufficient to allow utilization of l-glutamate as sole carbon source (107). Their nature and relative importance were clearly resolved for E. coli W in an analysis by Schellenberg and Furlong (244), using inhibitors and mutants that affect the individual systems. Some of their properties are described in Table 1, which was adapted mainly from Schellenberg and Furlong (244). Similar systems, cited by these authors, have been reported for other strains of E. coli and for S. typhimurium.
Table 1Properties of glutamate transport systems |
Halpern and his coworkers (107, 130, 166, 167) isolated structural and regulatory mutants for the l-glutamate-specific permease, GltS, and showed that it is active in membrane vesicles. GltS is a membrane protein of about 35,000 Da (58). The gltP gene of E. coli K-12, min 92.5 on the E. coli linkage map (153), was found to encode a polypeptide of about 38,000 Da, with four possible transmembrane segments (279). The E. coli l-glutamate-l-aspartate binding protein has a molecular weight of 31,000 and a Kd for glutamate of 0.7 μM (292). l-Glutamate- utilizing mutants of E. coli W and K-12 have severalfoldenhanced capacity for l-glutamate uptake via the l-glutamate-specific system (107, 166, 167). One class of mutations is closely linked to gltS (min 82 on the E. coli linkage map) and may be promoter types; the other class is unlinked (92 min) and probably affects a repressor gene for gltS. Kalman et al. (131) also characterized the gltS permease gene of E. coli K-12 and deduced a 42,000-Da product. Schellenberg and Furlong (244) subsequently showed that enhancement of expression of any of the three l-glutamate transport systems would allow utilization of exogenous l-glutamate by E. coli.
The regulation of synthesis of the l-glutamate permeases is not well understood. Kahane et al. (130) found gltS expression to be derepressed in growth of E. coli K-12 on l-aspartate as sole carbon or nitrogen source, but not when other carbon or nitrogen sources were present, suggesting that a metabolite of l-aspartate is an inducer. Kustu et al. (144) have shown synthesis of the l-glutamate-l-aspartate binding protein to be under nitrogen control in S. typhimurium, and Willis and Furlong (293) found the E. coli protein to be three- to fivefold repressed in growth on glucose as opposed to succinate.
L-Glutamine.
Wild-type E. coli K-12 can utilize l-glutamine as sole carbon and energy source, although growth is slow (171). Mutants showing enhanced transport grow well. There are several enzymes, glutaminases and glutamate synthase, which can convert l-glutamine to l-glutamate and NH3 and whose relative contributions have not been defined.
For the Enterobacteriaceae, glutaminases per se have been studied most intensively in E. coli, which has been reported to form two, A and B, whose physiological function is unclear (116, 223). l-Asparaginase II (see below) also has weak glutaminase activity. A number of amidotransferases, such as anthranilate synthetase, have glutaminase activity as part of their catalytic function and may contribute to l-glutamine catabolism (116). Glutamate synthase (α-ketoglutarate + l-glutamine → 2 l-glutamate), is essential for l-glutamate synthesis in the Enterobacteriaceae at low NH3 concentrations (269) and may be the most important factor in l-glutamine dissimilation.
Glutaminase A, purified and characterized by Hartman (116), is a protein of 110,000 Da (pH optimum of 3.5 to 5.5) composed of four identical subunits. The rate of synthesis is very low in cells growing rapidly on glucose, and on depletion of the energy source it increases rapidly in high NH3 (116, 275). Prusiner et al., however, found this increase to be inhibited by cAMP-CAP (224, 225).
Glutaminase B has been purified several thousand-fold and shown to be an acidic protein of about 90,000 Da, probably composed of four subunits, and with a pH optimum of 7 to 9 (223). Its activity is similar to that of glutaminase A in logarithmically growing cells, and its synthesis does not appear to be regulated. Prusiner and Stadtman (223, 226) have shown that futile cycling between glutaminase and glutamine synthetase is avoided by feedback regulation. To our knowledge, no structural genes for glutaminases A and B have been identified.
The biochemistry and regulation of glutamate synthase have been reviewed by Tyler (269). The enzymes of E. coli and K. aerogenes have been purified and characterized; they are flavoproteins with subunits of 53,000 and 135,000 (E. coli) or 175,000 (K. aerogenes). The structural genes (gltB and gltD) have been located at min 70 on the E. coli linkage map (214) and at min 69 on the S. typhimurium linkage map (90). Transcription of gltB absolutely requires Lrp (78). The role of this enzyme in l-glutamine catabolism and the nature of its regulation are discussed in more detail by Reitzer in this volume (chapter 23).
l-Glutamine transport has been characterized in E. coli (283, 284, 292) and S. typhimurium (17). Each organism possesses a high-affinity (Km, 0.1–0.2 μM) binding protein-dependent system and a low-affinity (Km, 10 μM) system.
The structural genes for the binding protein(glnP) and for at least the membrane component of the high-affinity system are located at min 17.7 on the E. coli linkage map (171). The glnP components of the S. typhimurium system are located at min 20 on the S. typhimurium linkage map (16). E. coli mutants selected by Masters and Hong (171) for rapid growth on l-glutamine as carbon source had up to sixfold-enhanced levels of l-glutamine transport. The mutations mapped at glnP. Mutants unable to form the glnP permease could not utilize l-glutamine as carbon source and invariably resulted in loss of ability of the cells to grow on l-glutamate or l-proline, although the parental strain grew fairly well on these compounds. No explanation was found for this second defect. glnP expression in E. coli and S. typhimurium is strongly repressed (up to 20-fold) when NH3 is readily available (17, 294). The E. coli glnHPQ region contains two promoters (208). One has a recognizable σ 70 initiation sequence and is apparently constitutively expressed. The other contains a consensus σ 54 initiation sequence and a consensus recognition site for the nitrogen regulator NR1. The latter promoter interacts with σ 54 polymerase to form a closed complex, and the transition to open complex is enhanced by NR1-phosphate and integration host factor (43). Gehring et al. (94) also observed a stimulation of l-glutamine transport in high sucrose, though not in high NaCl. The rationale for such a response may be the protective effect of l-glutamate in osmotic stress, which was discussed above.
Low-affinity l-glutamine transport most likely is by the osmotic shock-sensitive l-glutamate-l-aspartate transport system. Its binding protein has a low affinity for l-glutamine, and its capacity for l-glutamine uptake is suppressed in excess l-glutamate (284).
L-Arginine and Related Compounds.
L-Arginine can serve as sole carbon and nitrogen source for K. aerogenes, as a poor nitrogen source for S. typhimurium and E. coli, and as a source of carbon for some E. coli and Proteus strains (55, 88, 144, 251). L-Ornithine, an intermediate in L-arginine biosynthesis, is a precursor in putrescine biosynthesis, as is L-arginine for cells growing with excess L-arginine (55), and L-arginine is of course essential for protein synthesis. L-Arginine and L-ornithine are also substrates for biodegradative arginine and ornithine decarboxylases (192). In E. coli, formation of these latter enzymes is induced only at low pH in complex media (192). Their primary function is not entirely clear, but they are essential to growth under acidic conditions. They seem otherwise to be marginal to considerations of carbon flow and will not be considered further.
Friedrich et al. (87, 88) demonstrated the existence of an l-arginine utilization (aut) pathway in K. aerogenes, inducible by l-arginine, that can convert l-arginine to α-ketoglutarate via glutamate, allows utilization of l-arginine as sole carbon source, and does not generate urea. Cells grown with l-arginine accumulate some l-ornithine and form an acetylornithine 5-aminotransferase (Km, 1.1 mM; 59,000 Da; two subunits) capable of converting l-ornithine to glutamic semialdehyde, while cells grown on other nitrogen sources do neither. l-Arginine can support growth of an l-glutamate-requiring mutant. Evidence of Cunin et al. (55) suggests that the pathway proceeds via succinylated intermediates to glutamate and α-ketoglutarate, as in Pseudomonas cepacia (274). The pathway is subject to glucose repression, which is overridden by nitrogen control when l-arginine serves as nitrogen source.
E. coli strains generally do not use l-arginine as sole carbon and energy source. They do, however, have the genetic capacity to degrade it to succinate (55, 137, 251). The pathway (251, 297) consists of six enzymatic steps which convert l-arginine to succinate:
(1) l-Arginine → agmatine: arginine decarboxylase
(2) Agmatine → putrescine: agmatine ureohydrolase
(3) Putrescine → γ-aminobutyraldehyde: putrescine aminotransferase
(4) γ-Aminobutyraldehyde → γ-aminobutyrate: γ-aminobutyraldehyde transferase
The remainder of the pathway (steps 5 and 6), γ-aminobutyrate → → succinate (69), is described in the next section (γ-Aminobutyrate). The relevant E. coli K-12 genes and their map locations (9) are: reaction 1, speA, min 64; reaction 2, speB, min 64; reaction 3, pat, min 89; reaction 4, prr, min 31. l-Ornithine is also utilized via this pathway, by the reaction
l-Ornithine → putrescine: ornithine decarboxylase (speC, min 64)
Using a series of mutants blocked in various steps of this pathway, Shaibe et al. (251) demonstrated that E. coli K-12 can utilize it to convert arginine, agmatine, putrescine, and γ-aminobutyric acid to succinate. E. coli B also elaborates this pathway, and Kim (137) obtained mutants with enhanced pyridoxal phosphate-dependent putrescine transaminase levels which could use putrescine as sole carbon source.
Halpern and his coworkers (251, 252) found that the synthesis of arginine and ornithine decarboxylases was insensitive to glucose and to nitrogen control and that the synthesis of agmatine ureohydrolase was induced by l-arginine and agmatine. Induced synthesis was subject to about a 20-fold glucose repression, but the glucose effect was overridden by nitrogen limitation. There was little or no induction of putrescine aminotransferase or γ-aminobutyraldehyde dehydrogenase by l-arginine or putrescine, but synthesis of each enzyme was about 20-fold repressed in growth on glucose. The repression was not overcome by cAMP-CAP, but was lifted on nitrogen limitation. The two enzymes that convert γ-aminobutyrate to succinate are subject to glucose repression and nitrogen control, as described below. The decarboxylase results disagree with those of Boyle and his colleagues (23, 190, 191), who found synthesis of arginine and ornithine decarboxylases to be enhanced in growth on glucose, repressed at the level of transcription by cAMP-CAP, induced by their substrates, and repressed by putrescine. The disagreement appears to be due to strain differences (108). The decarboxylases are essential biosynthetic enzymes for putrescine synthesis and logically should not be subject to glucose repression. The other enzymes fill in the degradative pathway leading to succinate; carbon and nitrogen controls are appropriate for them and may allow scavenging of arginine and its derivatives when present in excess.
L. Reitzer (personal communication) has recently demonstrated the occurrence in E. coli of three enzymes of the l-arginine-l-glutamate pathway discussed above, which involves succinylated intermediates. Their expression is induced by nitrogen limitation and involves the Ntr system. Thus this pathway may also contribute to l-arginine dissimilation in E. coli.
The "biosynthetic" arginine decarboxylase, agmatine ureohydrolase, and ornithine decarboxylase of E. coli have been purified and characterized, as have the biodegradative decarboxylases (23, 190, 267). The biodegradative decarboxylases are found in only about 10% of E. coli strains (23). The aminotransferase and dehydrogenase are less well characterized.
K. aerogenes has only a very weak arginine decarboxylase activity, presumably just sufficient for putrescine synthesis as needed. However, agmatine can serve as carbon and energy source in this organism, via the putrescine-γ-aminobutyrate pathway as in E. coli (89). The first enzyme was induced by agmatine and the others by putrescine and γ-aminobutyric acid. All are subject to glucose repression, overridden on nitrogen limitation.
Kustu et al. (144) reported that S. typhimurium grows only very slowly with l-arginine as sole nitrogen source, but that it grows well with l-arginylarginine as nitrogen source and that mutants altered in a nitrogen control that affects several permeases also grow well. This finding suggests that S. typhimurium might possess the genetic capacity to utilize l-arginine as a carbon source.
l-Arginine transport in E. coli is mediated by two binding-protein-dependent systems: a high-affinity l-lysine-arginine-ornithine permease (Km for arginine, 5 × 10–9 M) and a lower-affinity l-arginine-ornithine permease (Km for arginine, 1.25 × 10–7 M; for l-ornithine, 10–6 M) (34, 239, 240). The relative importance of these two systems to l-arginine catabolism in E. coli has not been defined. The regulation of l-arginine transport is distinct from that of l-arginine biosynthesis, as mutants (argR) that affect expression of genes for biosynthesis do not affect expression of genes for transport and vice versa (33).
The binding protein for the 26,000-Da l-lysine-arginine-ornithine system of E. coli is the argT gene product, located at min 50 on the E. coli linkage map (91, 209, 239). Its expression is repressed by l-lysine (38). The binding protein for the 28,000- to 33,000-Da lower-affinity arginine-ornithine system of E. coli (91) is the abpS gene product, min 60 on the E. coli linkage map. Mutations in an adjacent regulatory site (abpR) result in enhanced expression of abpS (35). Substrate-binding protein interaction appears to be the rate-limiting step for transport by this system (35). Its synthesis is repressed by both l-arginine and l-ornithine (38).
Celis (36, 37) and Urban and Celis (272) have purified and characterized a kinase-ATPase of E. coli K-12 (argK gene product, min 63) that phosphorylates both of the periplasmic l-arginine binding proteins. An argK mutant showed reduced transport activity by both systems.
l-Arginine transport is not well characterized in S. typhimurium. Kustu et al. (144) showed that it severely limits l-arginine catabolism. Two uptake systems have been identified. One involves an l-lysine-arginine-ornithine binding protein, 26,000 Da, which functions through interaction with the hisP membrane complex (K m for l-arginine, 5 × 10–5 M) (91, 120, 143, 262). It is the product of the argT gene, at min 46 on the S. typhimurium linkage map. Its synthesis is under nitrogen and catabolite control. Mutations that release the nitrogen control enhance the formation of the binding protein and the utilization of l-arginine as nitrogen source in S. typhimurium (144). The other uptake system is a high-affinity l-arginine-specific system involving ArgP (Km for l-arginine, 3 × 10–8 M) (228, 239).
l-Arginine transport has not been characterized in K. aerogenes, but it is sufficient in the wild type to support l-arginine utilization as carbon source (269).
γ-Aminobutyrate.
γ-Aminobutyrate is the product of the glutamate decarboxylase reaction in E. coli (168) and also is an intermediate in the pathway of putrescine catabolism in E. coli and K. aerogenes (89, 251). It is likely that small amounts are present in the natural environments of the members of the Enterobacteriaceae, as a result of miscellaneous degradative processes.
Friedrich and Magasanik (89) have shown that γ-aminobutyrate can serve as sole carbon and nitrogen source in K. aerogenes and have characterized the pathway:
γ-Aminobutyrate + α-ketoglutarate → succinate semialdehyde + l-glutamate: γ-aminobutyrate aminotransferase
Succinate semialdehyde + NAD+ → succinate + NADH++H+: succinate semialdehyde dehydrogenase
The l-glutamate generated in the first reaction is recycled to α-ketoglutarate by the transaminase and lyase described under l-glutamate utilization (l-Glutamate, above). Synthesis of the two γ-aminobutyrate-degrading enzymes is induced by γ-aminobutyrate. It is subject to glucose repression, which is overridden by nitrogen control when nitrogen is limiting. To our knowledge, the K. aerogenes system has not been further investigated.
The situation in E. coli, mainly studied by Halpern and his colleagues, appears more complex, possibly because it has been examined in greater detail. The pathway gab is the same (67). It is not induced by γ-aminobutyrate, but is expressed at a low constitutive level, insufficient for the utilization of γ-aminobutyrate as sole carbon or nitrogen source (68). Mutants selected to use γ-aminobutyrate as nitrogen source contained eightfold higher levels of both enzymes. In secondary mutants selected to utilize γ-aminobutyrate as carbon source, uptake was enhanced. The system is subject to glucose repression and nitrogen control (181, 182, 307). Metzer and Halpern showed that formation of the γ-aminobutyrate system is repressed sevenfold in growth on glucose. The repression is lifted when γ-aminobutyrate is sole nitrogen source (Ntr control), but it is not reversed by cAMP (307).
Halpern and his coworkers (129, 183, 184) located the gab genes at min 58 on the E. coli linkage map, in the order gabP-(gabPp)-[gab(D,T)p]-gab(D,T)-gabC. The gabP gene encodes a specific transport system with a Km for γ-aminobutyrate of 12 μM. The permease is presumably a transmembrane protein. The gabT gene encodes the glutamic acid-succinic semialdehyde aminotransferase (Km, 1.3 × 10–3 M); the gabD gene encodes the succinic semialdehyde dehydrogenase (Km, 2.5 × 10–4 M); gabC is the locus, presumably regulatory, in which mutations allowing utilization of γ-aminobutyrate as carbon and nitrogen source occur. Because of apparent differences in sensitivity of gabD and gabT expression to glucose repression, Dover and Halpern (69) considered it possible that the genes have separate promoters, in spite of in vitro data. In vitro expression of gab genes cloned on appropriate restriction fragments yielded the following proteins: gabD, 54,000 Da; gabT, 45,000 to 48,000 Da; gabC, 40,000 to 43,000 Da. The gabC gene must have its own promoter, as a fragment carrying it alone was expressed, but the orientation is not known. The gabP gene product, a protein of 27,000 Da, was identified in an in vivo expression system. Whether the gabC gene product serves as an activator or repressor of gabDTP expression has not been determined.
Bartsch et al. (11) have also recently analyzed the gab gene cluster and have sequenced the gabT gene. The gene encodes a polypeptide of 46,000 Da. They found a gene order of gabP-(gabPp)-gabT-gabD-(gabT,D)p-gabC, differing from that of Metzer and Halpern (183) in the orientation of the gabTD genes. The reason for the discrepancy is not known.
L-Histidine.
L-Histidine is not utilized as a carbon and energy source by E. coli or wild-type S. typhimurium, but it is readily utilized via an inducible histidase (hut) system in K. aerogenes (156, 177). Magasanik and his coworkers showed that wild-type strains of S. typhimurium have the genetic potential to form this enzymatic sequence, but that it is not expressed sufficiently to allow the use of L-histidine as a carbon source (24, 156, 177). Regulatory mutants that can so utilize L-histidine have been isolated (see below). E. coli lacks the hut genes (97).
l-Histidine degradation in K. aerogenes proceeds through four enzymatic steps to l-glutamate and formamide, with NH4 + liberated in the first step (156, 157):
(1) L-Histidine → urocanate + NH4 +: L-histidine ammonia-lyase, or histidase
(2) Urocanate + H2O → 4-imidazolone-5-propionate: urocanase
(3) 4-Imidazolone-5-propionate + H2O → N-formimino-L-glutamate: 4-imidazolone-5-propionate amidohydrolase
(4) N-Formimino-L-glutamate + 2H2O → L - glutamate + HCONH2: N-formimino-L-glutamate formiminohydrolase
The actual inducer for the hut pathway is urocanate, the substrate for the second enzyme, rather than l-histidine (246). The primary role of the Hut enzymes appears to be carbon and energy generation, as opposed to nitrogen generation. The pathway is induced as a unit (245), although only the first enzyme, commonly referred to as histidase, is necessary to nitrogen generation. Moreover, synthesis of the Hut enzymes is subject to catabolite repression in both K. aerogenes and S. typhimurium. Neidhardt and Magasanik (197) observed, however, that if nitrogen availability is limited and l-histidine is available, formation of the Hut enzymes is induced in K. aerogenes regardless of whether conditions for catabolite repression exist or not. Bender (14, 15) and his collaborators have recently shown that this catabolite override is mediated by the product of the nac gene. Nac is a protein that couples transcription of several σ 70 operons, including put, that generate NH3, to the σ 54 Ntr nitrogen control. There is no such override in S. typhimurium, which lacks a functional nac gene, but E. coli is nac +. Nitrogen control of Hut gene expression is discussed by Reitzer in this volume (chapter 23) and so will not be considered here.
Only the third and fourth enzymes of the Hut pathway have been characterized in the Enterobacteriaceae. The imidazolepropionate hydrolase of S. typhimurium has a pH optimum of 7.4 and an apparent Km of 0.1 mM and was unstable in cell abstracts (256). Lund and Magasanik (154) showed N-formimino-l-glutamate formiminohydrolase of K. aerogenes to have a pH optimum of 8.5 to 9 and a Km of 4 × 10–2 M. The first two enzymes have been characterized by Phillips and his collaborators in Pseudomonas putida. The histidase is a 220,000-Da protein of four identical subunits (pH optimum, 9.0; Km, 5.3 mM) (46, 47). The urocanase is a protein of 110,000 Da, with one tightly bound NAD+ per molecule as cofactor, a pH optimum of 7 to 8, and a Km of 0.066 M (73, 74, 95).
l-Histidine transport has been studied most intensively in S. typhimurium by Ames and her collaborators. At least five systems are capable of l-histidine uptake in this organism. The most sensitive and specific l-histidine permease is the hisJQMP product (Km , about 10–8 M). It is a four-component system, involving a periplasmic binding protein (hisJ product) and three membrane proteins, which is under nitrogen control (144). The respective four genes constitute an operon at min 46 on the S. typhimurium linkage map (91, 121). An analogous system exists in E. coli; the genetic locus is at min 50. A second system, with a Km for l-histidine of about 10–6, involves the l-lysine-arginine-ornithine binding protein, the argT gene product, which is nitrogen and catabolite controlled (144, 262). It, like the hisJ protein, interacts with the hisQMP membrane components. The argT locus is adjacent to the hisJQMP operon in S. typhimurium and E. coli, but is not part of it (144, 207). Three low-affinity systems include the aroP permease (Km for l-histidine, 10–4 M) and two uncharacterized systems, X and Y, each with a Km for l-histidine of about 10–4 M (4). There are no data on the relative importance of these transport systems to the utilization of l-histidine as a carbon source. l-Histidine transport, however, does not seem to be a limiting factor in utilization, as all mutations that enhance utilization have been shown to affect expression of hut genes (see below). aroP expression is repressible by l-phenylalanine and l-tyrosine (243). Schlesinger et al. (246) demonstrated a separate urocanate permease in K. aerogenes, product of the uut gene, but no counterpart was found in S. typhimurium (24).
The genetic and physical organizations of the hut systems are virtually the same in K. aerogenes and S. typhumurium. The hut locus is between gal and bio in both organisms, at min 18 on the S. typhimurium linkage map. The locus consists of two operons in S. typhimurium. The gene order is hut(M)IGC(P)UH. hutH, hutU, hutI, and hutG are the structural genes for the four enzymes governing steps 1 through 4 of the Hut pathway. hutC is the repressor gene, which governs expression of the hut genes, and hutC mutations are cis and trans recessive to hut C + (19, 98, 256, 258). hutM is a control region which governs expression of the hutIGC operon from a promoter that is proximal to hutI; hutP is a control region that governs expression of the hutUH operon from a promoter proximal to hutU. hutQ, a site in hutP, is apparently the hutUH operator, as mutations in it result in partially constitutive expression of hutUH. hutM, hutP, and hutQ mutations are cis dominant and trans recessive to their wild-type counterparts and thus are DNA regulatory sites (258). Transcription is unidirectional (48). The genetic arrangement is the same in K. aerogenes. However, in that organism hutC has its own promoter (247) and there is probably yet another promoter for hutG, since insertion mutations in hutI allow some expression of hutG (248). Blumenberg and Magasanik (19) showed that there is significant homology between the hut DNA of S. typhimurium and K. aerogenes, highest among the structural genes. The K. aerogenes hutC sequence shows a potential promoter and ribosome binding site upstream of a coding region for a 27,218-Da protein (247). None of the other enteric hut structural genes has been sequenced, but Schwacha et al. (248) sized the gene products in minicells as follows: HutI, 44 kDa; HutG, 33 kDa; HutU, 57 kDa; HutH, 54 kDa.
Many strains of S. typhimurium fail to use l-histidine even as a nitrogen source. Meiss et al. (177) showed that in strain LT-2 this failure is due to weak expression of the hutUH promoter. Mutations hutP 0 → hutP + result in a more efficient promoter for histidase and urocanase, allowing utilization of l-histidine as nitrogen source. A second mutation, hutM + → hutM, in the hutIGC control region, is necessary to enhance promoter efficiency for hutIG such that l-histidine can also serve as carbon source. Brill and Magasanik (24) showed that some strains, such as 15-59, are naturally hutP +. Strain 15-59 can use l-histidine as nitrogen source, but a mutation at hutM is still required for its use as carbon source. Smith et al. (256) (Table 2) showed that expression of hutUH increases 50- to 100-fold on induction, whereas expression of hutIG increases only 3- to 4-fold. Growth with glucose reduces the induction ratio for hutUH 10- to 20-fold and that for hutIG 2- to 3-fold. Parada and Magasanik (215) observed that cAMP largely reverses glucose repression on expression of hutUH, but not at all on expression of hutIG. This finding suggests that catabolite repression on hutIG may involve factors other than the cAMP-CAP systems. Mutations in hutC result in constitutive expression of hutIGUH; certain mutations in the hutP region (referred to as hutR) result in partial constitutivity and partial escape of hutUH from glucose repression (96, 206, 256). Cooper and Tyler (48) measured hut-specific mRNA synthesis in S. typhimurium hut + and regulatory mutant strains and found it to correlate well with enzyme levels, indicating transcriptional regulation. This is consistent with the observation of Hagen and Magasanik (104) that purified HutC protein binds specifically to hut DNA and that the binding is released by urocanate.
Table 2Hut enzyme levels in wild-type and mutant strains of S. typhimurium 15-59a |
The regulatory pattern governing expression of S. typhimurium hut gene expression allows for protection of the l-histidine supply for biosynthetic purposes by at least two buffering mechanisms. The inducer for the hut pathway is urocanate, the product of the first degradative reaction, rather than l-histidine (246). Thus, given the low uninduced level of histidase, a significant amount of histidine must be present in order to give rise to sufficient urocanate to trigger induction. Also, since the hutC gene is part of the hutIGC operon, induction of this operon increases the repressor level and has a damping effect (257) on further induction.
Regulation of hut gene expression in K. aerogenes is very similar to that in S. typhimurium and in hut operons transferred to E. coli (97, 197, 215, 247), except for the absence of nitrogen control in S. typhimurium. Klebsiella and Salmonella repressors regulate each other’s operons quite well, though the homologous repressors have a significantly stronger effect on hutIG expression (96).
In NH3 excess, K. aerogenes hut + requires cAMP-CAP for hut expression (222). Nieuwkoop et al. (205, 206) identified a single cAMP-CAP-dependent, in vitro hutUH transcript. In the absence of cAMP-CAP an oppositely oriented transcript was observed, originating about 80 bp upstream of the hutUH transcript start, whose start site overlaps a cAMP-CAP consensus site. RNA polymerase bound preferentially to the upstream start site in the absence of cAMP-CAP and to the hutUH-specific site in its presence, suggesting that the function of cAMP-CAP was to channel RNA polymerase to the "correct" promoter. A detailed analysis of the kinetics of open complex formation at the two promoters by Osuna et al. (213), however, indicated that cAMP-CAP directly affects hutUH transcription initiation. The presence of the other promoter thus appears to be fortuitous.
L-Proline.
L-Proline is utilized as carbon and nitrogen source for strains of E. coli, S. typhimurium, and K. aerogenes and is also an osmoprotectant (40, 41, 52, 83, 159, 232). Enzymology, genetic organization, and regulation of the L-proline utilization pathway (put) are very similar in the three organisms.
l-Proline catabolism involves two enzymatic steps in E. coli, an oxidation to pyrroline 5-carboxylate followed by an NAD-linked dehydrogenation to form l-glutamate, the reverse of the biosynthetic sequence (83). Although Δ-pyrroline-5-carboxylate is also an intermediate in the l-proline biosynthetic pathway, a mutant for the biosynthetic reductase reaction converted l-proline to l-glutamate as readily as the wild type did. The two degradative activities were inseparable, which suggested that they might form a complex, facilitating the compartmentalization of catabolism and biosynthesis. Dendinger and Brill found an analogous pathway in S. typhimurium and devised enzymatic assays (62), and Chen and Maloy demonstrated the system in K. aerogenes (40). A single protein (the putA gene product) was shown to catalyze both reactions in each organism (40).
The putA gene product is a remarkable protein. It catalyzes both of the degradative reactions and also acts as a repressor of put operon transcription in the absence of l-proline (1, 64, 100, 159, 179, 180). It is thus capable of specific interactions with the two cofactors flavin adenine dinucleotide (FAD) and NAD, with l-proline as both substrate and inducer, with the cytoplasmic side of the inner membrane, with put operator sites on the DNA, and with itself.
Menzel and Roth (179) showed the S. typhimurium dehydrogenase reaction to have a Ping-Pong mechanism involving reduction of FAD followed by its reoxidation through the respiratory chain. They found a very high Km for l-proline, about 50 mM. Using a less stressful purification, Abrahamson et al. (1) obtained a Km of 2.5 mM for the E. coli enzyme. The subunit molecular weight of the S. typhimurium protein is 132,000 (178). There is some question as to whether it functions as a monomer, as a dimer, or both. Menzel and Roth (179) observed enzymatic activity of the Salmonella enzyme with both monomers and dimers and suggested that it may exist as a monomer in the membrane and a dimer in the cytoplasm. Brown and Wood (25) purified the E. coli enzyme and found it to act in vitro as a dimer of about 293,000 Da.
The membrane association of the PutA protein is not a tight one; the protein dissociates upon salt or detergent treatment. As the reaction sequence requires electron transport coupling, however, artificial acceptors must be present if the solubilized protein is to be active (1, 63, 100, 180, 232). Futile l-proline-l-glutamate cycling is apparently avoided by the high Km (2.5 mM) for the proline dehydrogenase activity, the obligatory coupling of the two degradative activities, and a lack of induction of the put genes unless the l-proline pool is above normal (76). Moreover, proline oxidase is inhibited in high osmolarity, when excess l-proline is osmoprotective. It is likely that proline binding sites on the protein for induction and for enzymatic activity are identical, as a common class of putA mutations that results in noninducibility also increases the enzymatic Km for l-proline (162).
As a transcriptional regulatory element, the PutA protein appears unique. It is unusually large for a DNA binding protein, though it contains a potential helix-turn-helix sequence (3). The evidence for its role as the specific put repressor is very strong. Mutations to put constitutivity map throughout the putA gene (180). The purified PutA protein bound specifically to put operator sites shown by mutational analysis to be involved in repression, and the DNA-protein complexes showed proline dehydrogenase activity (63). These binding sites were methylated in vivo in the presence of l-proline, but not in its absence, in wild-type cells; in constitutive mutants they were methylated in vivo in the presence and absence of l-proline. In vitro, reversal of DNA binding specifically required the presence of an electron acceptor. This and other evidence suggested that the inductive process increases PutA protein hydrophobicity and thus favors its association with the membrane (63, 83). The model for induction, developed by Maloy and his coworkers, proposes that in the absence of l-proline, the S. typhimurium PutA protein locates in the cytoplasm and binds to operator sites on put DNA, acting as a repressor. In the presence of l-proline and an appropriate electron acceptor the protein is reduced, its hydrophobicity increases, and it becomes a membrane-associated enzyme (63, 64, 162). This model is in accord with the data and seems a very neat arrangement. It also has the advantage of sparing membrane attachment sites when they are not needed. A similar pattern of DNA and membrane binding was described with the E. coli protein by Wood (299) and Brown and Wood (25).
l-Proline uptake in E. coli, S. typhimurium, and K. aerogenes is primarily via an inducible l-proline permease, the product of putP, a gene adjacent to putA (28, 40, 160, 180, 298). The permease is coinduced with the PutA protein, and therefore its function is presumably primarily degradative. It accounts for 80 to 90% of l-proline uptake in fully induced cells of E. coli. PutP is an integral membrane protein with 12 membrane-spanning α-helices, which functions by Na+-proline symport and is essential for the use of l-proline as carbon and nitrogen source. Its Km for l-proline in S. typhimurium is 2 μM. Wood and Zadworny (300) observed very rapid metabolism of proline upon uptake in E. coli which, in view of the high Km of the oxidase, led them to suggest that uptake and oxidation may be coupled.
At high osmolarity, l-proline has another important role in the cell, as an osmoprotectant. A neutral, highly soluble amino acid, and rather abundant in nature, it is ideal for this purpose. Thus, mutants of S. typhimurium that overproduce l-proline show enhanced osmotolerance in proportion to the increased intracellular level of the amino acid, and l-proline added to the medium has a similar effect. No other exogenously supplied amino acid is protective. The l-proline biosynthetic pathway is not derepressed in high osmolarity, but synthesis and activity of the proP permease and synthesis of the proU permease, which also transport the osmoprotectant glycine betaine, are enhanced (26, 27, 54). This allows the cell to garner any extracellular l-proline that is available. Also, in this circumstance, PutP function is inhibited (189).
PutP is a very useful system for dissection of the mechanism of membrane transport, as both substrate- and ion-binding mutants are readily selected (65, 195). The S. typhimurium mutations map in several small clusters in putP and may define the putP sites for substrate and ion binding and translocation. Ohsawa et al. have obtained similar mutations in E. coli and have shown that the E. coli permease is also a membrane protein spanning 12 α-helices (211).
The put loci of E. coli, S. typhimurium, and K. aerogenes consist of the genes putA and putP in each case, divergently transcribed from a central control region. The S. typhimurium and E. coli loci are located at min 22 on the respective linkage maps. The sequences of the put control regions of the three genera and the putA and putP genes of S. typhimurium and E. coli are highly conserved among the genera (40, 188, 196, 298; J. M. Wood, personal communication). Maloy and his coworkers have shown the putA and putP promoters to overlap in E. coli and S. typhimurium. The transcripts do not overlap, but are adjacent, in a regulatory region defined by cis-dominant constitutivity mutations (64, 105, 106). In K. aerogenes, however, the transcription starts are well separated (40).
Expression of both putA and putP is subject to catabolite repression in the three genera, and in each case the control region contains three potential CRP binding sites (40). The catabolite repression effect in S. typhimurium is about fivefold on expression of putA and about twofold on expression of putP and is reversed by cAMP in growth on glucose (161, 199). Hahn and Maloy (105) obtained catabolite repression-resistant mutants in S. typhimurium which mapped in the putAP promoter region, were cis-acting, and in most cases completely relieved catabolite repression for both genes. The E. coli and K. aerogenes put genes are subject to nitrogen control, presumably via the nac system as there are no NtrC binding sites in the intergenic regions (40, 222; L.-M. Chen, T. Goss, R. Bender, and S. Maloy, submitted for publication). S. typhimurium put genes are not subject to nitrogen control, and there are no Nac binding sites in the control region.
In a detailed DNA binding and deletion analysis of the S. typhimurium put control region, deSpicer et al. (64) demonstrated PutA protein binding to multiple sites, including the genetically defined operator between the putA and putP transcription starts. They also showed that the region contained areas of curvature and that deletion of individual binding and/or curvature sites had significant effects on maximal expression and induction ratio for both putA and putP. O’Brien et al. (210) identified two IHF (integration host factor) binding sites in the intergenic region and found a mild enhancing effect of IHF on repression of putA and putP. The authors suggest that put repression involves binding of the regulatory proteins to multiple sites in the large intergenic region to form a looped structure, facilitated by the native curvature and IHF-induced bends.
The catabolism of l-aspartate and l-asparagine in E. coli has the unique feature that expression of aspartase and of the high-affinity asparaginase II is activated by FNR (product of the fnr gene; 9) and is greatly enhanced in anaerobiosis (127). The immediate product of these reactions is fumarate (170). It has been suggested that the enzymes might function in anaerobiosis for the generation of fumarate as an electron acceptor (124), or that FNR was recruited as a regulatory element by these pathways because of an as yet undefined anaerobic function that they serve (I. R. Beacham, personal communication).
L-Aspartate.
L-Aspartate utilization has been studied mainly in E. coli, where Kay (132) has shown that it can serve as sole carbon source. S. typhimurium (144) and K. aerogenes (269) utilize it as sole nitrogen source, and since the aspartase reaction also generates fumarate, it must provide some carbon in those organisms. Aspartase functions anaerobically in E. coli via the aspartate aminotransferase and aspartase reactions to provide fumarate, an electron acceptor and precursor of succinate, from glucose (51). It is an essential component of the pathway of L-glutamate degradation in E. coli (170), as described above, and L-aspartate is a precursor of L-arginine (55).
Aspartase (l-aspartate ammonia-lyase) catalyzes the reaction (291):
l-Aspartate → fumarate + NH4 +
The E. coli enzyme (193,000 Da; four identical subunits) has been purified and characterized by Suzuki et al. and Guest et al. (102, 266). It appears to be a single species. The Km for the forward reaction is 10–3 M and the pH optimum is 8.7. It is a trans-elimination, analogous to the reactions catalyzed by an E. coli fumarase and argininosuccinase, enzymes with which aspartase has significant structural homology and likely a common evolutionary origin (302). The structural gene, aspA, of E. coli K-12 is located at min 94 on the E. coli linkage map, adjacent to the fumarate reductase gene frdA (169, 260). It consists of 1,431 bp (303) and yields a 52,190-Da gene product. Sequences with homology to the consensus transcription start occur 48 and 165 bp upstream of the translation start, and sequences with homology to the CAP and FNR recognition sites are centered 380 bp upstream of the translation start, on opposite strands of the DNA (301). Tosa et al. (268) have shown that intracellular aspartase is partly bound to particles or membranes and appears to be more stable when so bound than in the soluble fraction.
The level of aspartase in E. coli under aerobic conditions is low in l-glutamate-utilizing mutants growing with ammonia and with glucose as carbon source and is fivefold enhanced in growth with l-glutamate as sole carbon source (109, 171). Woods and Guest (301), however, have shown that basal aspartase synthesis is fivefold repressed in growth with glucose, and the glucose effect is probably mediated by cAMP-CAP. It is thus not clear whether the regulation of aspartase synthesis also involves induction by l-glutamate or l-aspartate. Under anaerobic conditions aspartase contributes to the biosynthesis of succinate in growth on sugars. It is probably essential for this purpose in malate dehydrogenase mutants, since together with aspartate aminotransferase, it allows the cells to bypass the block between oxalacetate and fumarate (51). Jerlstrom et al. (127) and Woods and Guest (301) have shown it to be under oxygen control by FNR (10- to 15-fold derepression under anaerobic conditions in fnr + cells).
l-Aspartate can be transported in E. coli W by four distinct systems (132, 244): (1) a binding protein-dependent l-glutamate-l-aspartate system (Km for l-aspartate, 5 × 10–7 M); (2) a binding protein-independent l-glutamate-l-aspartate system (Km for l-aspartate, 4 × 10–6 M;. (3) a binding protein-independent l-aspartate-specific system (Km, 3.7 × 10–6 M); and (4) a dicarboxylic acid system (succinate, malate, fumarate, and l-aspartate) (Km, 3 × 10–5 M). Schellenberg and Furlong (244), by means of strains mutant for each of these uptake systems, analyzed their contributions to L-aspartate uptake. In lactate-grown cells, system 1 contributed 25%; system 2, 52%; system 3, 23%; and system 4, a negligible amount.
Systems 1 and 2 of E. coli are discussed under L-glutamate catabolism (see L-Glutamate, above). Aksamit et al. (2) have purified a 30,000-Da l-aspartate-l-glutamate binding protein, similar in size to its E. coli counterpart (292), from S. typhimurium.
System 3 of E. coli, l-aspartate specific, accumulates l-aspartate in membrane vesicles and is therefore presumably a transmembrane system. It does not appear to be regulated by l-aspartate (132, 134). Aksamit et al. (2) also demonstrated an l-aspartate membrane transport system in S. typhimurium. Its activity is not inhibited by l-glutamate, so it is likely the counterpart of the third E. coli system.
System 4 of E. coli is a dicarboxylic acid transport system (dct), active on succinate, malate, and l-aspartate (132, 149, 237). Its synthesis is induced 25- to 30-fold by succinate and other non-amino dicarboxylic acids, but apparently not by l-aspartate, and the induction is blocked in growth on glucose (132, 134). The glucose effect is overcome by cAMP, and cya mutants are not inducible (149). Mutations in three loci, dctA (min 80 on the E. coli linkage map) and dctB and cbt (min 16 on the E. coli linkage map) (133, 149, 150), abolish growth on dicarboxylic acids, including l-aspartate, as carbon sources.
L-Asparagine.
The literature suggests that l-asparagine can be used anaerobically by wild-type E. coli as carbon source and aerobically as nitrogen source, but probably cannot be used aerobically as carbon source (295). l-Asparagine is converted to l-aspartic acid by l-asparaginase, which is synthesized by both S. typhimurium and E. coli. Whether l-asparagine can be used depends both on the entry mechanism, which is limiting in at least some strains of E. coli, and on the regulation of expression of the enzyme.
The l-asparaginases are among the most thoroughly studied E. coli enzymes because of the use of one of them, l-asparaginase II, in the treatment of acute lymphoblastic leukemia (249). E. coli synthesizes two quite different l-asparaginases (29). l-Asparaginase I, product of the ansA gene (min 39 on the E. coli linkage map) (52, 261), is a low-affinity cytoplasmic enzyme. l-Asparginase II, product of the ansB gene (min 63.8 on the E. coli linkage map) (20), has a higher affinity, is secreted into the periplasm, and is made only under anaerobic conditions (31). Both E. coli enzymes have been purified and characterized, and their genes have been sequenced (82, 126, 261). The high-affinity l-asparaginase II (Km, 1.15 × 10–5 M) consists of four identical 33,000-Da subunits (122, 126, 261). This enzyme also has activity against l-glutamine, about 4% of its activity against l-asparagine, and could therefore be responsible for some or all of the l-glutaminase activity reported in E. coli (122). The lower- affinity l-asparaginase I (Km, 3.5 × 10–3 M) has been less intensively studied. Asparaginases I and II are likely derived from a single ancestral gene, since they have two regions of strong similarity, with 24% sequence identity (115, 126).
Expression from E. coli asnB is regulated cooperatively by two global regulators, CAP and FNR (123, 124). The enzyme is induced 100- to 1,000-fold (32) in wild-type cells anaerobically, in the absence of glucose and other competing sugars. Strains lacking either regulator have very low levels of the enzyme. Expression from S. typhimurium asnB is positively regulated by CAP and anaerobiosis, but it is not regulated by FNR (125).
One metabolic role of l-asparaginase is to provide nitrogen and carbon from exogenously provided l-asparagine. This should be equally important aerobically as anaerobically, and yet l-asparaginase II is made mainly anaerobically. Jennings and Beacham (124) suggested that it has a different metabolic role, i.e., the provision of fumarate as a terminal electron acceptor. A cell provided with l-asparagine and glycerol could then grow anaerobically by coupling glycerol oxidation to fumarate reduction, which might explain why it is under FNR control. However, Beacham also suggests (personal communication) that the cell may have a need for l-asparaginase anaerobically for some other reason, and have recruited FNR regulation for this purpose.
The uptake of l-asparagine has been relatively little studied. In E. coli there appear to be at least two systems for l-asparagine entry, a high-affinity system repressed by growth with l-asparagine and needed for growth at lower asparagine concentrations, and a second, low-affinity system (295, 296).
In S. typhimurium, a gene (ansP) which encodes an l-asparagine permease has been cloned (M. P. Jennings, J. K. Anderson, and I. R. Beacham, submitted for publication). The deduced protein sequence showed significant homology with a family of basic and aromatic amino acid permeases, including AroP and PheP, and suggested the presence of 12 transmembrane domains (Jennings et al., submitted).
Tryptophanase and tyrosine phenol-lyase, the enzymes of l-tryptophan and l-tyrosine catabolism, are present in only a few genera of the Enterobacteriaceae, and both activities are not usually present in the same genus. However, there is clearly a close evolutionary relationship between them, as the respective proteins of E. coli and Citrobacter freundii are about 50% homologous (136). The E. coli tryptophanase and the Citrobacter tyrosine phenol-lyase have significant l-cysteine-degrading activity (142, 259), and the S. typhimurium cysteine desulfhydrase has tryptophanase activity (101, 139, 140), though l-cysteine does not induce the aromatic enzymes, nor tryptophan the cysteine enzyme. A rationale for such a relationship is not obvious.
L-Cysteine and L-Cystine.
L-Cysteine is quite toxic to enteric bacteria and is therefore utilized with care, as described below. Gutnick et al. (103) have shown that it can serve as sole carbon and nitrogen source in S. typhimurium. Tyler (269) reported that K. aerogenes can use it as a nitrogen source, thereby presumably deriving some carbon from it, but that E. coli does not utilize it.
l-Cysteine catabolism in S. typhimurium has been shown by Kredich et al. (139, 140) and Guarneros and Ortega (101) to be via a cysteine desulfhydrase that is induced up to 1,000-fold above basal level by l-cysteine or l-cystine. The rate of synthesis is not affected by glucose, indicating that this system is not under catabolite control, although it does provide carbon (101). Its primary function is probably detoxification. It may also provide S= for synthesis of other sulfur-containing compounds. Its specificity appears to be high, and it has no cystathionase or tryptophan synthetase activity and only a low tryptophanase activity. S. typhimurium does not use l-tryptophan as sole carbon or nitrogen source, but one may speculate that some l-tryptophan utilization might occur in the presence of l-cysteine.
Cysteine desulfhydrase is a hexamer of molecular weight 229,000 with one pyridoxal phosphate per monomer. Its Km for l-cysteine is 2.1 × 10–4 M, and its pH optimum is 8.5. The l-cysteine reaction has been proposed to proceed via an unstable intermediate, 2-aminoacrylate. This compound can react with a second l-cysteine to give 2-methyl-2,4-thiazolidine-dicarboxylate, or can decompose to pyruvate and ammonia. Less than stoichiometric amounts of pyruvate are therefore formed when purified enzyme is used for the reaction. Crude cellular preparations contain another factor which inhibits the 2-amino acrylate + l-cysteine condensation, so in the intact cell the reaction products are probably just pyruvate, ammonia, and H2S(139, 140).
Guarneros and Ortega (101) have demonstrated an l-cysteine desulfhydrase activity in E. coli, induced by l-cysteine or l-cystine. Its rate of synthesis is enhanced in growth on glucose, and its function is presumably detoxification and provision of S=. Its pH optimum is 7.2, significantly lower than that of the Salmonella enzyme.
The E. coli tryptophanase also catalyzes the reaction
l-Cysteine + H2O → pyruvate + NH3 + H2S
The relative activity of the crystalline enzyme is the same with either l-cysteine or l-tryptophan as substrate (204). Its synthesis is induced only by l-tryptophan, however, so it can contribute to l-cysteine utilization only when l-tryptophan is also present. We are not aware of any studies on l-cysteine catabolism in K. aerogenes.
There appear to be three l-cystine transport systems (10). The primary one, CTS-1, involves a binding protein and is responsible for most of the cystine uptake at concentrations above 20 μM. Its Km for l-cysteine is 2.0 × 10–6 M, and its V max is 9.5 nmol/min. Its expression is enhanced in sulfur limitation and is CysB dependent. The other two systems can transport sufficient external l-cystine to permit growth of a cys mutant. The second system, CTS-2, has a higher affinity (Km for l-cystine, 5 × 10–8 M) and a lower capacity (V max , 0.22 nmol/min). The third system is not well characterized. Expression of the second and third systems does not appear to be regulated.
L-Tryptophan.
Most strains of E. coli utilize l-tryptophan as carbon and nitrogen source via an inducible l-tryptophanase-l-tryptophan permease system that converts it to pyruvate, NH3, and indole (259). Some Shigella, Klebsiella, and Proteus strains degrade l-tryptophan similarly, but S. typhimurium, K. aerogenes, and most of the other Enterobacteriaceae do not (61). The tryptophanase system of E. coli K-12 is expressed to a high level in response to l-tryptophan (several hundred-fold induction), is extremely sensitive to catabolite repression, is not subject to nitrogen regulation, and is not subject to regulation by the trp operon repressor TrpR (114, 158, 259, 263). The control pattern clearly marks it as a generator of carbon and energy, yet one wonders why such a system should have developed in only a few enteric genera. l-Tryptophan in our experience does not support proficient growth of E. coli, but exposure to high levels of it were not deleterious, at least in the laboratory (61, 259; unpublished data), and apart from an inducible l-tyrosine phenol lyase in a few other genera (142), we are not aware of analogous activities for the other aromatic amino acids among the Enterobacteriaceae.
Newton and Snell (203, 204, 259) found the crystalline enzyme (tnaA gene product) to consist of four subunits, each with a pyridoxal phosphate cofactor linked to the ε NH2 of a lysine. It is a cytoplasmic protein with a relatively high Km for l-tryptophan of 0.33 mM and a pH optimum of 7.2 to 8.8. It catalyzes three classes of α,β elimination reactions:
l-Tryptophan + H2O → indole + pyruvate + NH3
l-Serine → pyruvate + NH3
l-Cysteine + H2O → pyruvate + NH3 + H2S
and two classes of β-replacement reactions:
Indole + l-serine → l-tryphophan + H2O
Indole + l-cysteine → l-tryptophan + H2S
The tryptophanase reaction is reversible (282) in high concentrations of pyruvate and NH3. Under these conditions, the enzyme can substitute for tryptophan synthetase in a trpAB mutant. A number of l-tryptophan analogs are substrates for tryptophanase. Phillips and his coworkers have utilized them to examine the reaction mechanism and sequence in detail and have used site-directed mutagenesis to examine the roles of specific amino acids in the reaction (217, 218, 220). They found that the indole NH is likely involved in substrate-enzyme interaction, as its modification is inhibitory. The reaction proceeds through two quinoid intermediates to the aminoacrylate with the release of indole, and the aminoacrylate is subsequently converted in several steps to the end products. A lysine at position 269 is probably involved with the quinonoid reaction steps, as the quinonoid portions of the sequence are inhibited when this lysine is replaced by arginine (220). The enzyme’s active site is located in or near a cysteine at position 298, as replacement of this amino acid results in altered cofactor affinity (218). This is likely the cysteine whose reaction with N-ethylmaleimide was observed by Raibaud and Goldberg (230) to inhibit activity. l-Tryptophan residues at positions 298 and 330 are not essential; they can be replaced by phenylalanine without affecting activity (219).
It is possible that a posttranslational step is involved in the formation of tryptophanase. Belezikian et al. (13) found a time lag for the appearance of active enzyme of 1 to 2 min after the completion of translation.
l-Tryptophan is transported by E. coli via three permease systems: the general aromatic amino acid transport system (aroP gene product; Km, 4 × 10–7 M), which also carries l-phenylalamine and l-tyrosine; mtr, an l-tryptophan-specific permease (Km, 10–6 M) which is also responsible for indole transport; and the l-tryptophan-specific tnaB gene product, a lower-affinity (Km, about 7 × 10–5 M), high-capacity carrier that is coinduced with tryptophanase and subject to catabolite repression. The relevant genes have been located at min 3, 69, and 83, respectively, on the E. coli map and sequenced, and the predicted protein structures suggest that each permease is a transmembrane protein (42, 72, 117, 198, 242, 243). The mtr and tnaB gene products show considerable homology to each other and to the tyrP gene, which codes for the l-tyrosine-specific permease. Sarsero and Pittard (242) found that these three proteins appear to have only 11 membrane-spanning domains, as opposed to the usual 12. They are, however, rather small, about 400 amino acids each, and have unique structural features that may be consistent with an 11-span arrangement (242, 243).
Yanofsky et al. (306) have shown that the TnaB permease is essential for utilization of l-tryptophan as carbon source in wild-type cells. Its primary function thus appears to be the scavenging of tryptophan as a carbon source, consistent with its regulatory pattern. The regulatory patterns for Mtr and AroP suggest that they are more concerned with provision of amino acids for protein synthesis (242).
One might expect that the induction of high levels of tryptophanase could deplete the l-trytophan pool sufficiently to be detrimental under certain conditions. This does not seem to be a significant problem, probably because of the high tryptophanase Km. Yanofsky et al. (306) have found that wild-type cells grown in a low-catabolite medium with l-tryptophan show only a mild and transient excess of anthranilate synthetase synthesis, as compared to an isogenic tnaA mutant, when they are transferred to the same medium lacking l-tryptophan.
The tnaAB genes constitute an operon that is located at min 83 of the E. coli K-12 linkage map (72). tnaA is the promoter-proximal gene; it specifies a 471-amino-acid polypeptide (56) that is identical in composition to the 471-amino-acid polypeptide of E. coli B (128). tnaB specifies a 415-amino-acid polypeptide in E. coli K-12 (242, 243). There is a 319-bp leader region between the transcription start and the tnaA translation start which is concerned with induction, which is actually the lifting of an attenuation control, in the presence of tryptophan (57, 263; chapter 81, this volume). Although constitutive with regard to this induction control, the tna promoter is highly sensitive to catabolite repression. Tryptophanase expression is reduced 100-fold in growth on glucose, as opposed to growth on glycerol. A CAP consensus site is located just upstream of the transcription start, and the cAMP-CAP complex is necessary to RNA polymerase binding at that site and to tna transcription initiation in vitro.
The results from a number of experiments have suggested that catabolite repression in the tna system involves more than the action of the cAMP-CAP complex at the promoter. This does not seem to be the case. rho mutations did not alter the catabolite control on tna expression, making unlikely a rho involvement. M. Eshoo and C. Yanofsky (unpublished data) have replaced the tna promoter with the tet promoter, leaving the leader region intact. In this situation, with tna expression driven by the tet promoter, there is no repression in growth on glucose. The experiment rules out a role for the leader region in the catabolite control and indicates that the entire glucose effect is at the cAMP-CAP-controlled promoter. Some of the confusion on this issue concerns the TnaΒ permease. A high concentration of l-tryptophan is required for induction (57), and when expression of the tna operon, including tnaB, is repressed by glucose, the level reached can be suboptimal for induction and the catabolite effect can thus appear larger than it should. In cells with high levels of the mtr permease, this complication is avoided (C. Yanofsky, unpublished data).
L-Tyrosine.
E. coli and Salmonella spp. do not utilize l-tyrosine as a carbon and energy source. However, Citrobacter (Escherichia) intermedius, C. freundii, and Erwinia herbicola strains (5, 142) all form an inducible l-tyrosine phenol-lyase, with C. intermedius able to grow slowly with l-tyrosine as sole carbon and nitrogen source. The distribution of tyrosine phenol-lyase among the enterobacterial genera is limited, and the physiological role is unclear.
Tyrosine phenol-lyase catalyzes the reaction
l-Tyrosine + H2O → phenol + pyruvate + NH3
The structural gene specifies a protein of 51,000 Da, with 456 amino acid residues (5). The enzyme, which comprises four identical subunits, has been purified and crystallized (5, 60). Each subunit binds one pyridoxal phosphate via a lysine residue. The pH optimum is about pH 8 and the Km for l-tyrosine is 0.2 mM. Tyrosine phenol-lyase is apparently specific to the l-isomer. It is closely related to the E. coli tryptophanase, with 43% identity in the amino acid sequence, and the chemical mechanisms of the reactions are very similar (136). It also converts l-cysteine and l-serine to pyruvate (142). The enzyme is inducible (142), and Phillips and his coworkers (R. S. Phillips, personal communication) have found that l-tyrosine enhances its level 25-fold in cultures of C. freundii. There is no evidence for a sequence encoding a leader peptide in the DNA preceding the structural gene (5, 136). The regulation thus apparently differs significantly from that of tryptophanase, in spite of the close structural relationship between the two enzymes.
In E. coli, as described in the tryptophan section, l-tyrosine is transported by a specific permease (tyrP gene product) and by the general aromatic amino acid transport system (aroP gene product) (42, 243), but we are not aware of any uptake studies in the tyrosine utilizers.
Members of the Enterobacteriaceae have the genetic potential to express at least two l-serine deaminases, two l-threonine-l-serine deaminases, and one l-threonine dehydrogenase (253, 265, 270). There is no evidence, however, that any of these activities is primarily directed to carbon utilization. Wild-type E. coli does not utilize either l-serine or l-threonine as carbon source, and the major l-serine deaminase and the threonine dehydrogenase are regulated by Lrp, not CAP (85, 148, 200). The physiological roles of this group of degradative pathways in wild-type cells are obscure.
L-Serine.
E. coli K-12 does not use l-serine as sole carbon and energy source unless it is also provided with glycine, l-leucine, l-isoleucine, and l-valine. E. coli K-12 also removes l-serine rapidly and earlier than any other amino acid, during growth in tryptone broth (227). It mutates readily to use l-serine (in the presence of l-isoleucine, to avoid l-serine toxicity) (273). S. typhimurium and K. aerogenes apparently differ from E. coli in that the wild-type strains do use l-serine in the absence of the other amino acids (103, 277). K. aerogenes grows slowly on l-serine and mutates readily to grow rapidly (277). Early references that suggest that E. coli does use l-serine are difficult to assess because of the then frequent use of dl-serine in assays.
l-Serine degradation involves only one enzymatic step, deamination of l-serine by l-serine deaminase, which converts l-serine to pyruvate and ammonia (216). E. coli K-12 synthesizes two such enzymes, l-serine deaminases 1 and 2, which are highly similar in sequence (73% identity) (265; Z. Q. Shao, R. T. Lin, and E. B. Newman, submitted for publication) and which consist of single polypeptide chains with a molecular weight of 49,000.
Their high structural similarity suggests that l-serine deaminases 1 and 2 are mechanistically similar. However, only l-serine deaminase 1 has been studied in detail (99). It has an extremely high Km for l-serine (10–2 M). Pyridoxal phosphate is not a cofactor, and no other cofactor has been found. Similar enzymes from anaerobic bacteria have been considered to be iron-sulfur proteins (99). The large number of cysteines in the E. coli molecule, as well as the activation by iron, suggest that the E. coli enzyme may also be an iron-sulfur protein.
The in vitro assay of l-serine deaminase 1 and l-serine deaminase 2 requires incubation with iron and dithiothreitol to activate the enzyme, a posttranslational mechanism which is relatively unusual in E. coli. In extracts of mutants that cannot convert the inactive precursor to active enzyme, activity can be obtained by incubation with iron and dithiothreitol (201). The mechanism of activation is not known. It has been suggested that the enzyme, as it is synthesized, folds into an inactive conformation and that it is activated by refolding to a different conformation (J. Moniakis, H. S. Su, and E. B. Newman, submitted for publication).
The structural genes for these enzymes, sdaA (39.5 min) and sdaB (60.1 min), have both been sequenced (253, 264). sdaB is the downstream gene of a two-gene operon of which the first gene, sdaC, is a putative serine transporter (Shao et al., submitted) with a high similarity to the tdcC gene, which codes for an l-threonine transporter (250).
Regulation of expression of both sdaA and sdaB is remarkably complex (E. B. Newman, unpublished data; Shao et al., submitted). The sdaCB operon is transcribed in rich media, where it constitutes about 50% of the l-serine deaminase activity, the rest coming from sdaA. The sdaCB operon requires cAMP and CAP for transcription. sdaC transcription is induced two- to threefold by l-leucine and Lrp together. l-Serine deaminase 2 is not found in minimal medium, apparently due to an inefficient ribosome binding site (253).
Transcription of sdaA is induced by the amino acids glycine and l-leucine, but not by l-serine (216). It is also induced anaerobically, at high temperature, and as a response to DNA-damaging agents. Mutations in the membrane-bound protein Cpx (Ssd) greatly increase sdaA transcription. Lrp represses sdaA transcription, an effect which is partially reversed by leucine (148, 200). l-Serine can consequently be used as a carbon source when l-isoleucine is provided by either ssd or lrp mutants, and by the wild type when glycine and l-leucine are also provided (273). However, since both l-serine and l-leucine are toxic and since l-isoleucine overcomes both toxicities, addition of l-isoleucine is also required for growth on serine (273).
The l-serine deaminase of K. aerogenes is induced by l-threonine, as well as by glycine and l-leucine, but not by l-serine (277). Its synthesis is subject to catabolite control via cAMP-CAP, but apparently not to nitrogen control.
l-Serine deaminase is highly specific for l-serine, but can deaminate l-threonine to a limited extent. Enzymes which are known as threonine deaminases are much less specific. The K. aerogenes isoleucine-inhibited l-threonine deaminase can serve physiologically as an l-serine deaminase as judged by the fact that some mutants which grew well on l-serine had increased l-threonine deaminase, while others had increased l-serine deaminase (277). Dual action against l-serine and l-threonine is particularly notable with the biodegradative l-threonine deaminase, which should be known as an l-hydroxyacid deaminase. In cells grown anaerobically with rich medium and without glucose, this tdcB-encoded l-threonine deaminase must be a major catalyst of l-serine deamination, along with the two l-serine deaminases.
The E. coli tryptophanase and the C. intermedius tyrosine phenol-lyase also deaminate l-serine (142, 259) (see above), as does tryptophan synthetase (204, 305).
Two l-serine transporters have been described in E. coli. The major l-serine transporter of E. coli is a relatively low-specificity l-threonine-sensitive permease, energized by Na+ and thus dependent on Na/H antiport (110). Two Na+/H+ antiporters, encoded by nhaA and nhaB, have been described (135). Mutants lacking the NhaB antiporter cannot transport l-serine even though the Na+/l-serine component and the second antiporter still function (111, 135). However, they produce small amounts of a much more specific proton-driven transporter, resistant to l-threonine (111). Mutants with enhanced levels of the H+/l-serine transporter can use it as the sole transporter for growth on l-serine (135). Cells which overexpress sdaC show l-threonine-insensitive transport, suggesting that sdaC codes for the second l-serine transporter (Shao et al., submitted). l-Serine transport by the second transport system is induced by l-leucine (Shao et al., submitted) as is sdaC transcription, so that apparently Lrp/leucine induces both transport and degradation.
Two other systems which transport l-serine have been described (44, 238). The finding that the Na+/H+ antiport mutants are so seriously impaired in l-serine transport suggests that these are not quantitatively important. However, results appear to vary among strains, so l-serine transport may merit study in several backgrounds (110).
Glycine.
Glycine cannot be used as sole carbon source by wild-type E. coli. However, a glycine utilization pathway can be established by two mutations, one increasing l-serine deamination and the other probably increasing l-serine hydroxymethyltransferase. The slow growth which results seems to involve conversion of glycine to l-serine and l-serine to pyruvate (Newman, unpublished data). Glycine is usually transported by the CycA permease (Km , 6 × 10–5 M)(147) (see d-Serine below), but in some strains it is transported only by diffusion (186).
L-Threonine.
L-Threonine does not serve as sole carbon source in wild-type E. coli, the only member of the Enterobacteriaceae in which threonine catabolism has been well characterized (202, 221). However, E. coli has the genetic capacity to express four pathways of L-threonine degradation, as follows. (i) Threonine deaminase, biosynthetic, catalyzes the formation of α-ketobutyrate, a precursor of isoleucine, which regulates its activity (270, 271). Because α-ketobutyrate is not used aerobically as a carbon source, this pathway is unlikely to be important in threonine degradation. (ii) Threonine deaminase, degradative, is an anaerobic enzyme which also catalyzes α-ketobutyrate formation (93, 271, 304). This enzyme is made only in the presence of L-threonine and at least three other amino acids and the absence of glucose and oxygen (75). (iii) Threonine aldolase, functions in Pseudomonas putida to convert L-threonine to acetaldehyde and glycine (193). There are conflicting reports as to the presence of this enzyme in E. coli. If it is present, its levels are apparently insignificant (202). (iv) The coupled threonine dehydrogenase:2-amino-3-ketobutyrate CoA ligase reactions convert L-threonine to glycine and acetyl coenzyme A (CoA) (39, 202, 221).
The use of l-threonine via the dehydrogenase and ligase was demonstrated by Newman and collaborators. The enzymes are present at low levels in wild-type cells and are increased during growth in the presence of l-leucine (213, 221). Mutants which make severalfold more of both enzymes are able to grow with threonine as carbon source (39, 85). Wild-type Serratia marcescens produces high levels of these enzymes and is able to grow readily with threonine as sole carbon and nitrogen source (138).
The biochemistry of the dehydrogenase pathway has been studied in detail by Dekker and his colleagues (8, 22, 77). The pathway is considered to be:
l-Threonine + NAD+ → 2-amino-3-ketobutyrate + NADH + H+: dehydrogenase
2-Amino-3-ketobutyrate + CoA → glycine + acetyl-CoA: ligase
2-Amino-3-ketobutyrate spontaneously decarboxylates to form aminoacetone, which is frequently mentioned in the early literature but is not part of the pathway. A tight channelling between the two enzymes is therefore likely.
Threonine dehydrogenase is a 148,000-Da protein of four identical subunits. The enzyme has a pH optimum of 10.3 and Km values for l-threonine and NAD+ of 1.43 mM and 0.19 mM, respectively. It is activated about 10-fold by Mn2+ and Cd2+. The active site involves a cysteine residue at position 38. The enzyme has an absolute requirement for Zn2+ which is extremely tightly bound. The region containing the active site is homologous with those of 25 other alcohol dehydrogenases, indicating that the enzyme is a member of the mammalian and microbial zinc-containing long-chain alcohol/polyol dehydrogenase family (8, 22, 77).
The 2-amino-3-ketobutyrate CoA ligase was first demonstrated in E. coli by Chan and Newman (39). It was shown by Mukherjee and Dekker to be a pyridoxal phosphate-dependent protein of 85,000 Da consisting of two identical subunits. The reaction is reversible and is studied in the reverse direction because the instability of 2-amino-3-ketobutyrate prevents an assay in the forward direction. As assayed in the reverse direction, the enzyme has a pH optimum of 7.5 and Km values for acetyl-CoA and glycine of 59 μMand 12 μM, respectively (194).
l-Threonine dehydrogenase and 2-amino-3-ketobutyrate ligase are encoded by the tdh and kbl genes, which constitute an operon (oriented Pkbl tdh) located at 81.2 min on the E. coli linkage map (7, 8). This operon is repressed by Lrp and induced by l-leucine. The pathway does not function to any physiologically significant extent in glucose minimal medium. When its level was increased 100-fold by two uncharacterized mutations, l-threonine could be used both as the source of glycine and as carbon source (85). Use of l-threonine as a carbon source was also established by a single IS3-mediated alteration of the promoter (6), resulting in increase of both activities.
Alterations in the level of expression of the tdh operon have profound consequences for both biosynthesis and energy production. Cells in which the dehydrogenase and ligase enzymes are expressed constitutively can derive glycine from endogenously synthesized l-threonine. In this background, the glyA mutation, which prevents conversion of glycine to l-threonine, does not make the cell auxotrophic, whereas glyA cells in wild-type background require glycine (39, 85). Even without mutation, wild-type cells grown with glucose, l-threonine, and several other amino acids can make their l-serine from l-threonine (235, 236). Moreover, the entire pathway is (to a limited extent) reversible, so that glycine can serve as a source of l-threonine (165).
It is not known whether use of l-threonine as carbon source requires increased l-threonine transport. l-Threonine enters the cell via an Na+-coupled cotransport system which carries both l-threonine and l-serine (110, 113) and via the livJ binding protein (113). Both this second transport system and the tdh operon are regulated by Lrp and l-leucine, which should coordinate transport and degradation.
Degradation of l-threonine is very different in anaerobic rich medium (without glucose) and in glucose minimal medium. In the former conditions, l-threonine is degraded by threonine deaminase, biodegradative, which produces α-ketobutyrate. Whether the α-ketobutyrate can be metabolized further in anaerobic medium without glucose has not been investigated (304), but it may act as an electron or amino acceptor (75, 270). Since Lrp production is greatly decreased in rich medium, neither threonine dehydrogenase nor the LivJ uptake system would be expected to be formed in large amounts, though this has not been examined. How l-threonine enters the cell in anaerobic rich medium has also not been studied, nor is it known whether tdh is expressed anaerobically.
l-Alanine alone among the "natural" l-amino acids is catabolized as the d-isomer, via a specific catabolic alanine racemase and a d-amino acid dehydrogenase of broad specificity. Significant amounts of other d-amino acids, especially d-serine, are likely to be present in the enterobacterial milieu as a result of nonenzymatic isomerization reactions (86). Except for d-serine, which is degraded by a specific deaminase, they are generally also substrates of the dehydrogenase. The existence of a cAMP-CAP control on expression of both enzymes suggests that the degradation of their substrates may contribute some carbon and energy. Possibly a specific enzyme has evolved for d-serine because of the particularly high lability of l-serine.
D- and L-Alanine and D-Amino Acid Dehydrogenase.
Both d- and l- alanine serve as carbon and energy sources for E. coli and S. typhimurium (103, 146). A single metabolic pathway is involved, in which l-alanine is converted to d-alanine by a specific racemase and the d-alanine is then converted to pyruvate and ammonia by a membrane-bound d-amino acid dehydrogenase of broad specificity (146, 233, 281, 290). This is the only case in which it has been shown that an l-amino acid must be converted to the d form for utilization as carbon source. Both activities are specifically induced by l-alanine. They are subject to cAMP-CAP-mediated catabolite regulation but not to nitrogen control (287, 288, 290; E. Daub, Ph.D. thesis, Massachusetts Institute of Technology, Cambridge, 1986). There are at least two factors in metabolism of the enteric bacteria which complicate this short pathway. First, d-alanine stands at a branch point. It is indeed a carbon and energy source, but a certain level is also essential for cell envelope biosynthesis. Provision must therefore be made for a biosynthetic d-alanine pool. Both E. coli and S. typhimurium form secondary, "biosynthetic" alanine racemases at a low constitutive rate, adequate for envelope biosynthesis, that presumably serve this purpose. The Km values of these enzymes for l-alanine are severalfold lower than those of the catabolic isozymes, in accord with their more essential function (79, 280, 287). Second, the d-amino acid dehydrogenases of E. coli and S. typhimurium oxidize several other d-amino acids as well as d-alanine, including d-phenylalanine, d-methionine, and d-asparagine (141, 212, 290). Membrane vesicles isolated from cells that were grown on dl-alanine engage in active transport of l-amino acids, and this transport is energetically coupled to d-amino acid oxidation. Respiratory uncouplers abolish the uptake. How important a role this system plays in amino acid transport is not clear, but the unique enzymological and regulatory arrangement suggests that it may be significant under certain circumstances.
The enzymology of the alanine racemase reaction has been studied most intensively in S. typhimurium. The enzymes have been purified, and the genes sequenced, by Walsh and Botstein and their collaborators (92, 212, 280, 281; Daub, Ph.D. thesis). The amino acid is bound to pyridoxal phosphate, the cofactor, and the reaction consists of proton removal followed by replacement to generate the optical isomer. The catabolic racemase, product of the dadB gene, is a 39,044-Da monomer with one pyridoxal phosphate bound to an ε-amino group of lysine. The Km values are 8.2 mM and 2.1 mM, respectively, for l- and d-alanine, and the pH optimum is 9 to 9.8 (280). It is a soluble enzyme. The catabolic alanine racemase of E. coli, product of the dadX gene, which has been sequenced (151), is reported to be a dimer of molecular weight about 100,000, with Km values for l- and d-alanine of 1.3 mM and 1.2 mM, respectively (147, 287). This enzyme may have some membrane association, as addition of either d- or l- alanine to membranes drives the dehydrogenase-linked uptake of amino acids (234). The biosynthetic alanine racemase of S. typhimurium, product of the alr gene, has also been purified; it is a monomer of molecular weight 40,000, with about 43% sequence homology to the catabolic isozyme and an identical pyridoxal phosphate binding sequence. The Km values for l- and d-alanine are 2.7 mM and 2.0 mM, respectively, and the pH optimum is 8.4. An analogous enzyme is formed by E. coli (92, 212, 287). Daub (Ph.D. thesis) suggests that the catabolic and biosynthetic enzymes may have evolved from a common ancestor, with the catabolic racemase developing a regulatory system appropriate to its primary function. These enzymes, like most alanine racemases, are highly sensitive to inhibition by d-cycloserine and related compounds and thus are potential antibiotic targets (147).
The d-amino acid dehydrogenases of E. coli and S. typhimurium are less well characterized. Olsiewski et al. (212) partially purified the E. coli enzyme and reported it to be a membrane-bound iron-sulfur flavoprotein coupled to the respiratory chain and consisting of two nonidentical subunits of 45,000 and 55,000 Da. The smaller subunit is the product of the dadA gene, which has been sequenced (151). Raunio et al. (233, 234), who characterized the enzyme in cell envelope preparations, reported a pH optimum of 8.3 to 8.4 for the reaction and a Km of 6.6 mM for d-alanine, and showed convincingly that l-alanine was not a substrate. This had been suggested earlier by Vyshepan et al. (278), who showed that inhibition of alanine racemase by cycloserine blocked oxidation of l-alanine but not d-alanine. Both Olsiewski et al. (212) and Raunio et al. (233) also found significant oxidation of the d-amino acids asparagine, aspartate, methionine, phenylalanine, and tyrosine, small amounts of which likely derive from environmental effects on protein degradation. Kuhn and Somerville (141) and Wild et al. (290) showed that the E. coli enzyme also has some specificity for d-tryptophan, as mutants using it as an l-tryptophan source were readily selected. The primary rationale for the enzyme’s broad specificity is likely to be detoxification, as d-methionine, d-phenylalanine, d-aspartate, and d-tryptophan can be incorporated into the E. coli cell envelope, with deleterious results (30). Wild et al. (290) reported that the d-amino acid dehydrogenase of S. typhimurium has a slightly different specificity, also oxidizing d-leucine and d-histidine. When the enzyme is present, its d-amino acid substrates likely provide some carbon and ammonia to the cells.
d-Alanine transport in E. coli (Km, 2 μM) is primarily via the osmotic shock-insensitive cycA system, which is also one of the l-alanine carriers (see d-Serine, below) (49, 229, 238). cycA mutants cannot use d-alanine as carbon source. Residual l-alanine transport is osmotic shock sensitive and subject to repression and attenuation control by l-leucine (113, 228). As l-alanine inhibits l-leucine and l-isoleucine transport by the LIV-1 system, which is osmotic shock sensitive and repressed in excess l-leucine, LIV-1 is probably the second l-alanine transport system (113).
It may be noted that l-alanine can serve as an amino donor for l-valine biosynthesis in the reversible transaminase C reaction, which generates pyruvate. Since synthesis of transaminase C is repressed by l-alanine, however, the metabolic role of the enzyme is presumably l-alanine biosynthesis, not catabolism (285).
The dad genes have been mapped in several laboratories, and the results are in some conflict. It seems clear that the catabolic racemase and at least the smaller d-amino acid dehydrogenase subunit genes are very closely linked and are coregulated in both E. coli and S. typhimurium (151, 281, 287). Wasserman et al. (280, 281) have presented convincing evidence for a gene order dadAB hemA at min 36 on the S. typhimurium linkage map, based on cotransduction of each locus with the same Tn10 insertion. This is in agreement with earlier work of Wild and Klopotowski (289). In E. coli, Wild et al. (287) have presented convincing evidence for a gene order fadR dadRAX hemA at the analogous site, min 26, based on deletion mapping and three-point crosses. Their finding agrees with earlier work of Kuhn and Somerville (141). Franklin et al. (84) have located a dad gene at min 1 on the E. coli linkage map, between ara and leu, also by well-designed three-point crosses, in agreement with earlier findings of Beelen et al. (12). Their data suggest that it encodes the larger dehydrogenase subunit. They and Beelen et al. (12, 84) have also located a positively acting dad regulatory gene, dadQ, at min 99. The biosynthetic alanine racemase gene has been located at min 91 on the S. typhimurium linkage map (79). What may be its E. coli counterpart has been mapped by Wijsman (286) to approximately min 93 on the E. coli linkage map.
Expression of the catabolic alanine racemase and the d-amino acid dehydrogenase genes is coordinately regulated by both induction and catabolite repression in E. coli and S. typhimurium (281, 287; Daub, Ph.D. thesis). Mutations that result in an enhanced basal level of the dehydrogenase, and in insensitivity of its synthesis to catabolite repression, were obtained by selection for ability to utilize d-tryptophan as a source of l-tryptophan (141, 289). The mutations, dadR, map immediately upstream of dadA. In E. coli the dadAX cluster appears to constitute an operon (151, 287), but the Salmonella dadB gene has its own promoter (S. A. Wasserman, Ph.D. thesis, Massachusetts Institute of Technology, Cambridge, 1983).
A detailed examination of dad regulation was carried out in S. typhimurium by Daub, using dadB::Mud lac fusions (Daub, Ph.D. thesis). Her results are similar to and extend those of other workers who have examined regulation of synthesis of the catabolic racemase and the dehydrogenase in E. coli and S. typhimurium. Daub found that glucose-grown wild-type cells and cya and crp mutants contained a significant basal level of the catabolic alanine racemase. Wild-type cells growing in glycerol or l-alanine as carbon sources showed, respectively, twofold- and sixfold-higher levels of enzyme. Starvation for d-alanine enhanced expression threefold, suggesting the presence of a residual control for possible biosynthetic purposes. The induction and catabolite repression ratios for the dad system are rather small, perhaps reflecting a necessity for protection of the alanine pool and for a detoxifying role of the dehydrogenase. Surprisingly, no evidence for a nitrogen control was found, in spite of the fact that d-alanine is a good nitrogen source. There is no evidence for regulation of expression of the alr gene.
It is likely that the dad system is under positive control. Wild and Klopotowski (288) were unable to obtain constitutive mutants in S. typhimurium, and Franklin et al. (84) found only low constitutive mutants in E. coli. The latter group also determined that a mutation to noninducibility of the dehydrogenase was recessive to its wild-type allele, indicating that presence of a regulatory gene product is necessary for induction.
D-Serine.
D-Serine serves as a readily utilizable nitrogen source and poor carbon source for about 70% of the ECOR collection of E. coli strains, including most of the human strains (103, 155a, 269) and for strains of S. typhimurium and K. aerogenes. It is also a growth inhibitor in minimal media (173). It is converted to pyruvate and ammonia by the inducible enzyme D-serine deaminase (D-serine dehydratase, EC 4.2.1.14, dsdA gene product, min 51 on the E. coli linkage map).The dsd locus of E. coli K-12 has been sequenced (164; M. Nørregaard-Madsen, E. McFall, and P. Valentin-Hansen, submitted for publication). It comprises the dsdA gene, a specific regulatory gene dsdC, and a gene of unknown function dsdX, in the order dsdA-dsdX-pXA-pC-dsdC. The D-serine deaminase system has been characterized only in E. coli K-12, and therefore discussion will be limited to that organism.
The enzymatic reaction worked out by Snell and Shafer and their collaborators is complex (70, 71, 80, 81, 163, 185). It involves substrate binding to the pyridoxal phosphate cofactor, followed by a dehydration step to form the aminoacrylate, subsequent rearrangement and rehydration, and finally release of NH3 and pyruvate. The enzyme is highly specific to d-serine (Km , 0.32 mM) and d-threonine (Km , 3.2 mM), and there are no other substrates of significance. The dsdA gene encodes a polypeptide of 47,000 Da (164). The enzyme functions as a monomer and thus has been a useful model for pyridoxal phosphate-mediated reactions.
d-Serine is transported by the glycine-d-alanine-d-cycloserine permease (Km , 10–5 M), a membrane transport system thatis the product of the cycC gene (min 95.5) (49, 152, 229, 238). Its synthesis is unaffected by d-serine or its other substrates (49), but it may be regulated by Lrp, as growth in l-leucine, the Lrp effector, enhances glycine uptake (229, 238).
d-serine deaminase induction is mediated by a specific activator protein, the dsdC gene product, with d-serine (Km , ca. 7 × 10–3 mM) and d-threonine (Km , ca. 4 × 10–2 mM) as the only known effectors (118). The dsdC gene is adjacent to dsdA (173). d-Serine’s relatively low Km for induction as compared to that for the deaminase reaction means that induction can be effectively triggered by low concentrations of d-serine. This may be important under physiological conditions. d-Serine inhibits growth of E. coli K-12 by blocking l-serine and pantothenate biosynthesis; thus expression of the deaminase is essential for detoxification when the supply of these compounds is limited (50).
dsdA gene expression is tightly controlled. The basal level of d-serine deaminase in dsdC + and in dsdC::Mu strains is only about 35 molecules per cell (175). DsdC is absolutely required for its activation in vivo and in vitro (119, 175). In dsdC + strains containing sufficient cAMP-CAP, exposure to d-serine evokes a 3,000- to 5,000- fold increase in the rate of d-serine deaminase synthesis and a 500- to 1,000-fold increase in the absence of cAMP-CAP (172). DsdC represses its own synthesis fivefold in the absence of d-serine (175). These effects are transcriptional (119). Thus, the dsd system has a simple and efficient control pattern to handle both its catabolic and detoxification functions. The expression of dsdA is not subject to control by the Ntr system, ppGpp, attenuation, Lrp, or IHF, nor is dsdC expression regulated by cAMP-CAP (21, 173, 175; unpublished data).
dsdC – mutants are noninducible, as would be expected with a positively controlled system(18). dsdC(Con) (constitutive) mutants are usually low constitutive and partially trans dominant in F' merodiploids, also consistent with positive control (173). dsdC(Con) mutants require cAMP-CAP for constitutivity, suggesting that in this system CAP enhances DsdC activation of dsdA expression (173, 176). dsdAp(Con) (promoter constitutive) mutants are readily isolated as d-serine deaminase-forming revertants of dsdC – types(18). Most are fully constitutive, and all are independent of cAMP-CAP to the extent of the constitutivity.
Expression of dsdA + is affected by context and by novobiocin, implying a role for supercoiling (176). Present evidence indicates that the mechanism of dsdA activation is complex, involving interactions between DsdC, d-serine, cAMP-CAP, RNA polymerase, and DNA (173, 174, 176).
d-Serine is not a naturally occurring amino acid, and it has been a mystery as to why many of the enterics possess a highly specific enzyme with a several thousand-fold induction ratio (see below) to convert it to pyruvate. d-Serine is highly toxic, but what would be the source of sufficient d-serine to cause a problem? Friedman (86) has shown that even at moderate temperatures there is significant nonenzymatic racemization of some amino acids, most particularly of l-serine. Thus sufficient amounts of d-serine to provide carbon and/or to cause toxicity may be generated from l-serine in the biosphere, and this may be a selective factor for the existence of a specific deaminase.
Neither E. coli nor S. typhimurium secretes proteinases, and therefore neither organism degrades extracellular protein (187), though other enteric bacteria do (e.g., Erwinia) (145). Both, however, make a variety of proteinases active against endogenously synthesized intracellular proteins. These bacteria grow very rapidly at the expense of protein hydrolysates used in a variety of natural media, e.g., tryptone or casein hydrolysates which contain both amino acids and oligopeptides. Bacteria in nature are more likely to encounter amino acids as hydrolysates of proteins than as single amino acids and thus must have evolved to handle mixtures of amino acids and oligopeptides. However, we found only one study on mixtures (227).
Pruss et al. studied the order in which amino acids disappeared from cultures of E. coli in tryptone broth and found that their strain removed l-serine first, and very rapidly, certainly without time to select a mutant (227). Utilization of l-aspartate followed shortly after but was slow; l-tryptophan, l-glutamate, l-alanine, and l-threonine were all used rapidly, but only after about a 4-h lag; and glycine was used slightly later and slowly.
This is reasonably close to what one might expect from the work on individual amino acids described above. l-Serine could be used if glycine, l-leucine, and l-isoleucine were present. The other amino acids known to serve separately as carbon sources were degraded. The relatively rapid use of glycine seems surprising, since 50% of the threonine carbon metabolized by cells growing with l-threonine as carbon source is converted to glycine and excreted (39). On the other hand, l-alanine is used rapidly as sole carbon source so that its slow use in mixtures is unexpected. This may reflect difficulties of l-alanine uptake from amino acid mixtures. l-Alanine enters on the same system that handles d-serine and glycine, both of which may compete with it for entry (238).
Further studies of this kind, using mixtures of amino acids in varying proportions and checking the cells for induction of transport mechanisms and of degrading enzymes, would give us a much clearer idea as to what cells do in nature. A comparison with different E. coli strains, as well as the other enterics, would also be useful.
Growth on peptides is a special case of the use of hydrolysates. Whether a peptide can be used depends on whether it can be transported and digested to single amino acids and whether the constituent amino acids can be used as an energy source. The enterics make a variety of dipeptide and oligopeptide transporters and a startling number of peptidases, of which Miller has described several in both E. coli and S. typhimurium without apparently coming close to enumerating all of them. Among those recently described is dipeptidyl carboxypeptidase, a C-terminal Zn2+ exopeptidase seen in both E. coli and S. typhimurium (45, 112). The Salmonella gene (dcp) has been cloned and sequenced.
Peptidases are important both for the complete degradation of intracellular protein and for the use of externally provided oligopeptides. The subject of peptide transport and degradation has been reviewed by Miller (187; chapter 62).
As we have seen, few amino acids support rapid growth in wild-type strains of E. coli and S. typhimurium when used as sole carbon sources. K. aerogenes is more proficient, growing well with l-histidine, l-proline, l-arginine, l-aspartate, l-asparagine, and l-glutamate (277). Yet in general, the Enterobacteriaceae grow very well with proteinaceous digests such as Casamino Acids.
There are several factors known to limit growth with individual amino acids, as follows. (i) One limiting factor is inadequate transport. Uptake severely limits the utilization of l-glutamate, l-glutamine, and l-cysteine in spite of the existence of multiple transport systems (10, 167, 171). In the case of l-cysteine this limitation is essential because of toxicity, but the rationale in the other two cases is not obvious. (ii) Degradative pathways are poorly expressed or cryptic. Thus, l-histidine is not used as sole carbon source by S. typhimurium, nor is l-arginine used by E. coli, because the respective pathways are not sufficiently expressed (19, 251). Again, the rationale is not obvious, as mutant strains with elevated enzyme levels can utilize these compounds. (iii) Degradative and/or uptake pathway expression depends on the presence or absence of secondary factors (Lrp, FNR). Thus, expression of l-serine deaminase, l-threonine dehydrogenase, and several other systems is regulated by Lrp/leucine (78, 148, 229, 238; Shao et al., submitted). Again, the rationale is not obvious, the metabolic role of Lrp remains unclear, and its role in transport regulation is not understood.
The family Enterobacteriaceae as a group grow well on mixtures of amino acids, which is presumably what they see in nature. Mixtures may have a number of interesting effects. l-Leucine and glycine modulate repression by Lrp, and so affect expression of several pathways (148). The presence of certain amino acids (i.e., l-isoleucine) prevents toxicity by others (i.e., l-serine) (273). Many permeases, such as CycC (d- and l-alanine, d-serine, glycine) (238), transport several amino acids, and competition for entry via these uptake systems may allow appropriate buffering of what enters the intracellular pools, thus preventing toxicities. And even though some of the degradative and transport systems may be poorly expressed in mixtures, the total effort may be adequate for growth. It is surprising that so little is known about the web of induction patterns and toxicities among amino acids, which affects expression of genes and cellular metabolism concerned with amino acid degradation in E. coli and S. typhimurium.
Amino acid biosynthesis is expensive for the cell. It is therefore essential that individual amino acids not be catabolized unless they are present in excess of the expected cellular requirements. The requirement of amino acids for protein synthesis is obvious. However, amino acids are used in other ways: for cell wall synthesis (e.g., d-alanine), for nitrogen transfer (l-glutamate, l-glutamine), for putrescine synthesis (l-arginine), or for osmotic protection (l-proline). Catabolism must be balanced against these requirements.
A variety of techniques are used for channelling amino acid towards and away from catabolism. Probably of primary importance are the generally high Km values of the degradative enzymes, which ensure that pools are not excessively depleted. Many catabolic pathways, as we have seen, are regulated by substrate induction, often coupled with catabolite repression and nitrogen control. These mechanisms assure that pathways are not expressed unless they can be useful. In some cases there are multiple enzymes catalyzing the same reaction which are differentially regulated, e.g., the biosynthetic and biodegradative alanine racemases (281). A particularly elegant arrangement exists for l-proline degradation (63). This pathway is the reverse of the biosynthetic pathway, but both degradative activities are present on the same polypeptide. The pool is protected by the high Km for l-proline of the first reaction and by the necessary coupling of the two degradative reactions.
There are a large number of issues that are presently not resolved. For example, most transport systems which have been studied are regulated by end-product repression, nitrogen control, or Lrp, but only a few respond to substrate induction and/or catabolite repression. Several degradative enzyme systems that generate carbon and energy are also Lrp regulated. As discussed above, the rationale for this control is unclear. The metabolic significance of a posttranslational mechanism that activates l-serine deaminase (201) is not understood, nor are the physiological roles of the hydroxyamino acid-degrading enzymes.
The channelling of amino acids is not perfect, and some exogenously provided amino acids are degraded even in growth with glucose and excess ammonium sulfate. This is true for l-serine: the l-serine requirement of a serA mutant was significantly decreased in E. coli upon introduction of a secondary mutation that eliminated l-serine deaminase (231). Similarly, in an l-proline-overproducing mutant of S. typhimurium, the intracellular l-proline pool increased threefold on introduction of a putA mutation (52). It is not known whether this is a more general phenomenon. The induction of tryptophanase (the only other system in which, to our knowledge, the question has been addressed) does not seem to have a significant effect on the l-tryptophan pool (306). Cells must of course degrade certain amino acids, even those that they synthesize endogenously, in order to form others (i.e., l-serine → glycine; l-threonine → l-isoleucine). These reactions are closely regulated, and whether other endogenously synthesized amino acids are attacked unnecessarily has not been investigated.
Finally, we must point out that research on amino acid catabolism has focused primarily on the fate of individual amino acids that were required to serve as sole carbon sources—and thereby, for practical purposes, as sole energy sources—in pure cultures. Pathways and control patterns have been defined on this basis. In the biosphere, however, these cells are presented with mixtures of amino acids, sugars, organic acids, etc., generally at lower concentrations than they see in the laboratory, and they usually must compete with other microorganisms for such nutrients. Interactions between amino acids are particularly likely to affect regulatory patterns and uptake patterns. Moreover, growth of cells for long periods under laboratory conditions would be expected to result in genotypic adaptations. It is possible that ancestral organisms may have been able to express some or all of the degradative pathways that are available to our current strains only as a result of mutations, and that such mutations simply reestablish ancestral characters. The laboratory situation, therefore, must be considered at best an approximation of the natural state. A comparison of patterns of amino acid utilization between defined laboratory strains and freshly isolated counterparts from the biosphere would be of considerable interest.
We are very grateful to I. R. Beachem, R. A. Bender, D. Botstein, R. T. F. Celis, E. Daub, M. Friedman, C. E. Furlong, J. R. Guest, Y. S. Halpern, N. M. Kredich, S. Kustu, W. K. Maas, B. Magasanik, S. R. Maloy, A. T. Phillips, R. S. Phillips, L. Reitzer, J. M. Wood, and C. Yanofsky for many helpful discussions, for reprints and preprints, and for critical reading of portions of the manuscript. This work was supported by American Cancer Society Grant no. NP-557 to E.M. and by Natural Sciences and Engineering Research Council (Canada) Grant no. OGP 000 6050 and a Killam Research Fellowship (Canada Council) to E.B.N.
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