Catabolism of Hexuronides, Hexuronates, Aldonates, and Aldarates

M.-A. Mandrand-Berthelot*, G. Condemine, and N. Hugouvieux-Cotte-Pattat

[SECTION EDITOR, AUGUST BÖCK]

Posted July 6, 2004

Unité de Microbiologie et Génétique, Centre National de la Recherche Scientifique / Université Claude Bernard Lyon 1 / Institut National des Sciences Appliquées, Domaine Scientifique de la Doua, 10, rue Raphaël Dubois, 69622 Villeurbanne, France

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Dedicated to the memory of Professor François Stoeber, deceased August 2003

Following elucidation of the regulation of the lactose operon in Escherichia coli, studies on the metabolism of many sugars were initiated in the early 1960s. The catabolic pathways of d-gluconate and of the two hexuronates, d-glucuronate and d-galacturonate, were investigated. The post genomic era has renewed interest in the study of these sugar acids and allowed the complete characterization of the d-gluconate pathway and the discovery of the catabolic pathways for l-idonate, d-glucarate, galactarate, and ketogluconates.

Among the various sugar acids that are utilized as sole carbon and energy sources to support growth of E. coli, galacturonate, glucuronate, and gluconate were shown to play an important role in the colonization of the mammalian large intestine. The biochemical reactions involved in the breakdown of sugar acids are limited to a few enzymatic activities, including mostly dehydratases, oxidoreductases, isomerases, kinases, and aldolases. The uptake of extracellular compounds is performed by specific transport systems. Most sugar acid transporters belong to the MFS and IT superfamilies. This clearly contrasts with hexose uptake, which is mainly mediated by PTS transport systems. Whatever the sugar acid used as substrate, the various routes lead to either the 6-carbon intermediate 2-keto-3-deoxy-6-phosphogluconate (KDPG) or the 3-carbon intermediates pyruvate and 3-phosphoglyceraldehyde. These compounds are further metabolized by the glycolytic pathway and provide energy via substrate-level phosphorylation. Pathway-specific transcriptional regulators allow the bacteria to adapt their catabolism to the carbon source(s) present in their environment. In the case of sugar acid degradation, the regulators often mediate negative control and are inactivated by interaction with a specific inducer, which is either the substrate or an intermediate of the catabolism. These regulators coordinate the synthesis of all the proteins involved in the same pathway and, in some cases, exert cross-pathway control between related catabolic pathways. This is particularly well illustrated in the case of hexuronide and hexuronate catabolism.

HEXURONIDES

Naturally occurring sugar acids derived from nutrients ingested by the host are present in the mammalian large intestine (135). In vertebrate liver, one of the major pathways of detoxification of endogenous and xenobiotic organic compounds is by conjugation of these aglycones to glucuronic acid. β-d-Glucuronide compounds are then excreted in the bile and provide a carbon source that can be efficiently exploited by intestinal Escherichia coli (36).

Hexuronides are hydrolyzed into hexuronates which are further metabolized, through the pathway discovered by Ashwell (2) and extensively studied by Stoeber and coworkers (138), into the two central metabolites pyruvate and 3-phosphoglyceraldehyde.

Transport and Hydrolysis of Hexuronides

The first enzyme of the hexuronide-hexuronate pathway, β-glucuronidase (β-glucuronoside glucuronosohydrolase, EC 3.2.1.31) (Table 1) (Fig. 1), was initially characterized by F. Stoeber, working on a fecal E. coli isolate, strain ML30 (137). He also demonstrated the existence of an inducible glucuronide-specific permease able actively to concentrate the substrate over 200-fold inside the cell. A wide variety of β-glucuronides are taken up and are able to induce β-glucuronidase activity; methyl-β-d-glucuronide is among the most effective (137). The compounds 5-bromo-4-chloro-3-indolyl-β-d-glucuronide, p-nitrophenyl-β-d-glucuronide (PNPG), and 4-methylumbelliferyl-β-d-glucuronide also act as powerful inducers (76). Subsequent investigations of the hexuronide-hexuronate pathways were conducted on E. coli K-12, which was more readily amenable to genetic analysis. In contrast to the fecal isolate, E. coli K-12 transports hexuronides poorly and shows only very low levels of β-glucuronidase induction. To achieve maximal levels of induction, external concentrations of inducer (10 mM) are required (137, 145).

Fig. 1Catabolic pathways for sugar acids in E. coli

Table 1Metabolic genes and enzymes involved in sugar acid pathways of E. coli

Both β-d-glucuronides and β-d-galacturonides are substrates of β-d-glucuronidase (34). In contrast to β-d-glucuronides, β-d-galacturonides cannot induce the synthesis of this enzyme (7) and thus cannot be used as sole carbon source by the wild-type strain. Methyl-β-galacturonide, therefore, has been used to select regulatory mutants constitutive for β-glucuronidase synthesis (91).

β-Glucuronidase was purified either by conventional methods (64, 67, 88, 137) or by affinity chromatography on a Sepharose column bound to the high-affinity competitive inhibitor thiophenyl-glucuronide (16). The enzyme is composed of four 68.2-kDa subunits, is resistant to thermal inactivation at 50°C, and exhibits a neutral pH optimum (34, 63, 88). The apparent K _m for PNPG was found to be 0.2 mM (67, 88). d-Glucaro-1,4-lactone, which is in equilibrium with d-glucarate, is a reversible competitive inhibitor of β-glucuronidase.

β-Glucuronidase has become most popular due to the extraordinary development of the GUS system in the late 1980s (42). Since β-glucuronidase is not widely distributed in eukarya and bacteria and because of the facile enzyme assay, the uidA (also called gusA) gene has been extensively used as a powerful reporter gene for analysis of gene expression in bacteria, animals, and plants. A great variety of gene fusion vectors, promoter-probe cassettes, and transposons have been developed (5, 6, 42, 65, 80).

Regulation of the uidABC Operon

The gene encoding β-glucuronidase, uidA, maps at min 36.5 on the E. coli chromosome (90), and it is transcribed counterclockwise (18) (Fig. 2). Two genes are cotranscribed with uidA (Fig. 2). The uidB (gusB) gene overlaps uidA by four base pairs. It is likely to direct the synthesis of the glucuronide permease, a protein with 12 membrane-spanning domains that bears similarity to the sodium:galactoside melibiose transporter (76) and belongs to the glycoside-pentoside-hexuronide (GPH):cation symporter family (TC #2.A.2.3.1) (Table 2). The uidC (gusC) gene encodes a predicted membrane-associated protein whose precise function in glucuronide metabolism remains unknown (145).

Fig. 2Genetic organization of sugar acid operons with their cognate regulatory circuits. Numbers refer to the position (min) of genes on the chromosome. The direction of the genes (represented by large arrows) shows their orientation on the bacterial chromosome (either clockwise or counterclockwise). The thin arrows represent the transcriptional effects of the regulators, with plain lines and broken lines to indicate primary and secondary levels of control, respectively. The + and – signs indicate positive and negative controls, respectively.

Table 2Transport systems involved in sugar acid pathways of E. coli

Directly upstream of uidA is the uidR (gusR) gene, which encodes a repressor specific for the uid operon (92). The uidA and uidR genes are not cotranscribed (15). Recessive mutations in uidR result in constitutive β-glucuronidase activity (92). Use of gene fusions demonstrated that uidR is negatively autoregulated and is sensitive to catabolite repression (15). Isolation and characterization of dominant-negative mutants indicated that the UidR protein acts as a multimer (19). UidR belongs to the TetR/AccR family of transcriptional regulators (Table 3). In addition to the regulation exerted by UidR, β-glucuronidase synthesis was found to be weakly repressed by the product of the uxuR gene, which primarily controls the uxuAB operon involved in the metabolism of d-glucuronate (93) (Fig. 2). In agreement with a dual repressive control mechanism by UidR and UxuR, full induction of β-glucuronidase synthesis depends upon the cooperative action of both β-d-glucuronide (or d-glucuronate) to relieve repression by UidR and d-fructuronate to relieve repression by UxuR (78, 89, 93). Moreover, the two gratuitous (nonmetabolized) inducers thiophenyl-β-d-glucuronide (137) and mannonic amide (7), which are structural analogues of β-d-glucuronide and d-fructuronate, respectively, act in concert to allow maximal induction of β-glucuronidase synthesis (78). When overproduced from multicopy plasmids, UidR and UxuR repressors are partially interchangeable for the control of uidA expression (116). Total repression of uidA requires the presence of both repressors, and, accordingly, two classes of presumptive operator-constitutive mutants affecting the binding of either UidR or UxuR have been described (93).

Table 3Regulatory genes controlling sugar acid metabolism in E. coli

The location of the transcription start point of uidA was determined by nuclease S1 mapping experiments (11). Analysis of uidA'-lacZ fusions suggested the presence of two putative operator sites allowing the binding of the two repressors UidR and UxuR (11). A recent comparative genomic approach has identified potential UxuR/ExuR binding sites upstream of both uidA and uidR (132), but the binding site for UidR remains to be established.

β-Glucuronidase synthesis is very sensitive to catabolite repression in the presence of glucose. This repression is reversed by exogenous addition of cyclic AMP (cAMP), and uidA is not expressed in a crp mutant (88). However, in vitro gel retardation and DNase I experiments failed to demonstrate specific binding of cAMP-cAMP receptor protein (CRP) in the regulatory region of uidA (11). The recent prediction of a relatively well-conserved CRP binding site upstream of uidA (132) invites the reexamination, by additional in vitro studies, of the mechanism whereby CRP and cAMP operate to activate uidA expression.

Occurrence of β-Glucuronidase Activity in Other Bacteria

The pathway for the utilization of glucuronides is unique to E. coli in sequenced gamma purple bacteria (132). Indeed, β-glucuronidase assays are routinely used as diagnostics for the specific detection of E. coli and Shigella species in clinical and environmental samples (25). Nevertheless, β-glucuronidase is found in other nonenterobacterial, anaerobic residents of the gut, particularly in the gram-positive genera Bacteroides and Clostridium, as well as in Streptococcus, Staphylococcus, and Corynebacterium spp. (145). The recent identification of the gusA gene of Lactobacillus gasseri ADH is the first report of a β-glucuronidase gene cloned from a bacterial source other than E. coli (39% identity at the amino acid level) (134).

HEXURONATES

d-Galacturonate is an important plant component since it is the main constituent of pectin. Free galacturonate can be liberated by the microbial flora in the intestine of herbivores. d-Glucuronate is found only at a low level in plant tissues. It is mainly found in animal mucus and in the form of β-glucuronides in the digestive tract. E. coli is able to use the two aldohexuronates d-galacturonate and d-glucuronate as sources of carbon and energy. Evidence of this catabolic pathway was first provided by Ashwell et al. (2). Later, Stoeber et al. (138) described more precisely the physiological and genetic aspects of the hexuronate pathway. After entering the cells by the ExuT transporter, d-galacturonate and d-glucuronate are degraded via parallel pathways involving consecutive isomerization, reduction, and dehydration steps (Fig. 1). d-Glucuronate can also be liberated intracellularly from β-d-glucuronides by the action of the β-glucuronidase UidA. The first enzyme common to both pathways, UxaC, isomerizes d-galacturonate and d-glucuronate into d-tagaturonate and d-fructuronate, respectively. d-Tagaturonate is then reduced by UxaB to d-altronate, which is dehydrated by UxaA. In the parallel pathway, d-fructuronate is reduced by UxuB to d-mannonate, which is dehydrated by UxuA. The two hexuronate pathways lead to the formation of a common intermediate, 2-keto-3-deoxy-d-gluconate (KDG). The first two intermediates of the hexuronate pathway, d-tagaturonate and d-fructuronate, can also support E. coli growth. In contrast, d-altronate and d-mannonate are not catabolized, probably because they cannot be taken up by the bacteria.

Biochemical Characterization of Hexuronate Catabolism

Each step of the two parallel galacturonate and glucuronate pathways was precisely characterized (Table 1). Active transport and accumulation of both hexuronates, d-galacturonate and d-glucuronate, is mediated by ExuT (66, 87). The fact that exuT mutants remain able to grow on d-fructuronate and d-tagaturonate indicates that these intermediates use separate transport systems to enter the cells (87). The apparent K _m for d-glucuronate uptake is 0.1 mM (Table 4). ExuT is a member of the major facilitator superfamily (MFS, TC #2.A.1) and probably functions as a hexuronate/proton symporter (Table 2). The hexuronate isomerase UxaC (EC 5.3.1.12) reversibly converts d-galacturonate and d-glucuronate to d-tagaturonate and d-fructuronate, respectively (3, 94, 144). Apparent K _ms of UxaC for d-galacturonate and d-glucuronate are 1.65 and 3.7 mM, respectively (Table 4). UxaC is also able to catalyze the reversible isomerization of d-tagaturonate into d-galacturonate with a twofold higher affinity for d-tagaturonate than for d-galacturonate. In contrast, the isomerization of d-glucuronate to d-fructuronate is nor reversible. The altronate oxidoreductase UxaB (EC 1.1.1.58) has strict substrate specificity towards d-tagaturonate with an optimal activity at pH 6.3 (47, 100, 106). The reaction is reversible, and the K _ms for d-tagaturonate and d-altronate are 350 μM and 90 μM, respectively. UxaB uses NADH but also NADPH as a cofactor, with K _ms of 60 μM and 400 μM, respectively. The mannonate oxidoreductase UxuB (EC 1.1.1.57) has a high affinity towards d-fructuronate (K _m of 0.5 mM) and NADH (K _m of 0.03 mM) and only a low affinity for d-tagaturonate (47, 101, 107). Apparent affinities for the substrates of the reverse reaction (d-mannonate and NAD+) are lower (Table 4). The altronate dehydratase UxaA (EC 4.2.1.7) is specific for d-altronate and has a K _m of 15 mM (122, 123, 136). Optimal activity of UxaA is at pH around 8 in the presence of Fe²⁺ ions and of a reducing agent such as β-mercaptoethanol. It is also activated by Fe³⁺ or Mn²⁺ ions and is inhibited by some acid sugars, mainly d-arabonate. Both Fe²⁺ and Mn²⁺ have a synergistic effect on UxaA activity, raising the possibility that UxaA has two binding sites for metal cations (35). The mannonate dehydratase UxuA (EC 4.2.1.8) is highly specific for d-mannonate, with a K _m of 20 mM (122, 123, 129) (Table 4). It has an optimal activity at pH 8.3 in the presence of β-mercaptoethanol and Fe²⁺. It is also activated by Fe³⁺ or Mn²⁺ ions and by Co²⁺, Ni²⁺, and Cu²⁺, but the latter function with less efficiency. There is a competitive inhibition of mannonate dehydration by d-sorbitol, d-mannitol, and d-gluconate. In contrast, most intermediates of the hexuronate pathway, namely, d-galacturonate, d-glucuronate, d-tagaturonate, and d-altronate, enhance activity. Upon Fe³⁺ activation, both UxaA and UxuA incorporate a single loosely bound Fe atom (with K_ds about 20 mM and 4.5 mM, respectively), which is not in a Fe-S core as observed for other Fe-hydratases (35). The reactions catalyzed by either UxaA or UxuA are not reversible, and these enzymes have no activity with KDG as substrate.

Table 4Biochemical properties of the proteins involved in hexuronate catabolism

Organization of Genes Involved in Hexuronate Catabolism

The structural genes encoding the different steps of hexuronate catabolism were identified by analysis of numerous mutants affected for growth with galacturonate or glucuronate (Table 1). Mutations in each step of the pathway were obtained by chemical mutagenesis and by insertion of Mu phages (52, 54, 55, 56, 87, 103, 125, 126). A specific method based on the toxic properties of an intermediate of the KDG catabolism, 6-phospho-KDG, was frequently used for isolation of mutants (110). Whereas an eda mutant grown in the presence of a hexuronate is poisoned by the accumulation of 6-phospho-KDG, a second mutation upstream in the pathway allows the eda mutant to grow in the presence of either d-galacturonate or d-glucuronate or both. This method was used to isolate several mutations in the hexuronate pathway. Localization of the mutations revealed that they are scattered in three regions of the E. coli chromosome (Fig. 2). The genes involved in d-galacturonate catabolism are located in two clusters, with the uxaB gene at min 34.6 (102) and the three genes exuT, uxaC, and uxaA at min 69.9 (87, 102, 103). The two genes, uxuA and uxuB, involved in the metabolic branch converting d-fructuronate to KDG are situated at min 98 (126). Coordinated synthesis of the corresponding enzymes suggested the existence of the two polycistronic transcriptional units uxaC-uxaA and uxuA-uxuB. This organization was confirmed by isolation of polar insertions in each gene of the hexuronate pathway. The phage Mud(Ap lac) was used to construct these polar insertions since they also gave rise to lacZ transcriptional fusions. Moreover, the transcriptional orientation of the genes relative to the E. coli chromosome was deduced by determination of the orientation of the Mu insertion in the transcriptional fusions (52, 54, 55, 56).

Regulation of Hexuronate Catabolism

The synthesis of the proteins involved in the hexuronate pathway is inducible (7, 105, 127). The ExuT transport system is synthesized at a high basal level, allowing the substrates to enter the cells and to induce the catabolic pathway. Growth in the presence of d-glucuronate leads to the induction of the transport system and of all the enzymes of the hexuronate pathway, including UxaA and UxaB, which are specific to d-galacturonate catabolism. In contrast, growth in the presence of d-galacturonate leads to the induction of only the transport system and the enzymes of the d-galacturonate pathway. Thus, UxuA and UxuB are induced only in the presence of d-glucuronate.

Isolation of mutants in which some of the catabolic steps are derepressed led to the identification of two negative regulatory genes located in the previously identified clusters. The exuR gene is in the vicinity of the exuT and uxaCA operons (104), and uxuR is near the uxuAB operon (128) (Fig. 2). The ExuR regulator governs expression of the genes involved in d-galacturonate catabolism, exuT, uxaB, and uxaCA, including the two steps common to d-galacturonate and d-glucuronate catabolism, i.e., their transport and isomerization. Analysis of mutations in each step indicated that the true inducers of the ExuR regulon are d-galacturonate as well as the first two intermediates of the hexuronate pathways, d-tagaturonate and d-fructuronate. For example, in an uxaC mutant, d-galacturonate is still able to induce ExuT and UxaB synthesis, while d-glucuronate has lost its inducing capacity. UxuR is the main regulator of the uxuAB operon encoding the two steps specific to d-glucuronate catabolism. The UxuR inducer is the first intermediate of the d-glucuronate pathway, d-fructuronate. In fact, the regulation of the uxuAB operon is more complex since it is mediated by the cooperative action of both ExuR and UxuR (128). UxuR also exerts a partial control on the β-glucuronide degradation, in concert with UidR (93).

The double control of hexuronate catabolism by ExuR and UxuR allows induction of the common steps, exuT and uxaC, in the presence of either d-galacturonate or d-glucuronate, as well as the repression of the glucuronate metabolism genes in the presence of d-galacturonate only. The ExuR-controlled operons (exuT, uxaCA, and uxaB) are derepressed by d-galacturonate or d-tagaturonate, but the glucuronate-specific operon (uxuAB) is still repressed in the presence of d-galacturonate. In the presence of d-glucuronate, d-fructuronate causes derepression of genes controlled by ExuR and of those controlled by UxuR. It is not possible to analyze the inducing capacities of altronate and mannonate, as they are not transported into E. coli cells. However, a structural analogue of mannonate, mannonic amide, is a gratuitous inducer of genes regulated by either ExuR or UxuR (7). The expression of all the genes of the hexuronate pathway, including the regulatory genes, is sensitive to catabolite repression exerted by glucose (52, 53, 56, 84, 87, 119). Their transcription is activated by the cAMP-CRP complex. Indeed, a typical CRP binding site is observed upstream of the exuT, uxaCA, uxaB, uxuAB, and uxuR operons (132).

Both ExuR and UxuR are negatively autoregulated (53, 56, 118, 119), with a stronger effect on ExuR than on UxuR (Fig. 2). Such an autoregulation in a negatively regulated system could allow a rapid adaptation to nutritional conditions. The catabolite repression of the regulatory genes could also participate in this effect. In the presence of a readily utilizable carbon source, expression of the regulatory genes is low but sufficient to repress the catabolic pathway, since the structural genes also need cAMP-CRP activation. The consequence of variations in the repressor concentration is that the control is highly dependent on the inducer concentration. For example, the inducer concentration sufficient to turn on the catabolic pathway is lower (in conditions of low repressor concentration) than the threshold concentration below which the pathway is turned off (in conditions of high repressor concentration). ExuR and UxuR are two regulators of the GntR family (Table 3). They share similarity both at the structural (49% identity) and functional levels. Increase in the ExuR and UxuR concentrations by the use of high-copy-number plasmids demonstrated that these regulators are capable of cross talk (116, 117). ExuR could replace UxuR for the regulation of the uxuAB and uxuR operons (116). In contrast, UxuR could not replace ExuR for the control of the exuT, uxaCA, or uxaB operons. Thus, ExuR and UxuR are only partially interchangeable.

Cloning of the exuT-uxaCA region was facilitated by the existence of a secondary λ attachment site in the regulatory gene exuR. An abnormal excision gave rise to a λ transducing phage containing the exuT-uxaCA genes (81, 83, 121). The other two regions, uxuAB-uxuR and uxaB, were isolated by complementation of mutations (14, 120) using the E. coli gene library of Clarke and Carbon (24). The corresponding regulatory regions were then analyzed by DNase I protection using RNA polymerase for uxaB and uxuAB (12, 17) and the cAMP-CRP complex for uxaCA (13). Isolation of an ExuR-LacZ hybrid protein showed that the N-terminal portion of ExuR is sufficient for its repressor activity in an in vitro transcription-translation system (82). Despite these advances, the DNA sequences of the ExuR or UxuR binding sites have yet to be defined. More recently, analysis of the ExuR-UxuR regulons by comparative genomics (132) showed that all the regulated operons are preceded by a palindromic sequence (AAATTGGTATACCAATTT) predicted to be the binding site of these regulators. Comparative genomic analyses failed to reveal differences between the sites which would allow discrimination between ExuR and UxuR. Even if they are capable of cross talk, these two regulators clearly have different specificities in vivo, which are probably reflected by minor variations in the binding sites leading to different levels of affinity for each operator. For example, increasing the copy number of the operator sites using multicopy plasmids showed that the affinity of ExuR for its operators varied. A higher affinity for the uxaB operator than for either the uxaCA or exuT operator was observed (14). The high affinity of UxuR for the uxuAB regulatory region is probably due to the presence of two binding sites in the regulatory region (17). The uidAB operon also possesses a potential UxuR binding site. Other genes were predicted to be members of the ExuR-UxuR regulons (132), including the uidR regulatory gene controlling β-glucuronide catabolism and the gntP gene encoding a gluconate transporter. Thus, the regulation of hexuronide-hexuronate catabolism involves the three regulators UidR, ExuR, and UxuR, whose syntheses are interconnected at the transcriptional level. These regulators are also likely to function in concert, and direct protein-protein interactions probably occur when they bind to adjacent or overlapping operator sites of the controlled genes.

Hexuronate Catabolism in Other Enterobacteria

Analysis of the Salmonella enterica serovar Typhimurium genome revealed that it has no uxaA or uxaB genes for the utilization of d-galacturonate. It also lacks the regulatory gene exuR. The S. enterica serovar Typhimurium genome, however, has the exuT gene and a putative uxuAB uxaC operon, as well as the regulatory gene uxuR, potentially allowing this bacterium to catabolize d-glucuronate (132). Early studies indicated that some Salmonella strains are able to use d-galacturonate as a carbon source. These strains were isolated from cold-blooded animals and from the environment, whereas galacturonate-nonutilizing Salmonella strains are most often isolated from humans and warm-blooded animals (75).

Yersinia pestis has orthologues of all the genes of the E. coli hexuronate catabolism, but the operon structures are different, with divergent uxuA and uxuB genes and a clustering of the genes involved in galacturonate catabolism, forming the uxaCBA operon (132).

To sustain its growth, Erwinia chrysanthemi can use pectin, oligogalacturonides, d-galacturonate, and its dehydrated derivative 5-keto-4-deoxyuronate (DKI) as carbon and energy sources. Since pectin degradation is a determinant of the pathogenicity of this bacterial species, the hexuronate pathway has been extensively studied in E. chrysanthemi (50, 57). The four steps involved in galacturonate assimilation are the same as those described in E. coli. They are encoded by two divergent operons, exuT and uxaCBA, controlled by the repressor ExuR. Thus, the operon structure is similar to that observed in Y. pestis. However, in contrast to E. coli, wild-type E. chrysanthemi strains do not assimilate β-d-glucuronides or d-glucuronate.

2-KETO-3-DEOXY-d-GLUCONATE

The two parallel pathways of degradation of d-glucuronate and d-galacturonate converge to 2-keto-3-deoxy-d-gluconate (KDG) (Fig.1). KDG is phosphorylated by the ATP-dependent KDG kinase, KdgK, to form 2-keto-3-deoxy-6-phosphogluconate (KDPG). KDG is a metabolite common to the Entner-Doudoroff pathway that drives catabolism of d-gluconate (38, 96). KDPG is then cleaved by the KDPG aldolase KdgA (also known as Entner-Doudoroff aldolase Eda), giving pyruvate and 3-phosphoglyceraldehyde. The triose-phosphate intermediate is further metabolized by the glycolytic pathway and provides energy via substrate-level phosphorylation. Mutants lacking KdgK show residual growth on either hexuronate, suggesting the existence of a secondary pathway from KDG (115). Enzymatic oxidation of KDG in the presence of NAD⁺ was demonstrated to occur (46). This reaction could be catalyzed by the oxidoreductase KduD, whose gene is widely distributed among the enterobacteria (Fig.1). The other steps of this secondary pathway are still unknown.

KDG can serve as the sole source of carbon and energy for E. coli K-12 only if synthesis of the KDG uptake system KdgT, which is strongly repressed and only weakly inducible in strain K-12 (113), is rendered constitutive by mutation (109, 115). Thus, the physiological role of KdgT could be the uptake of KDG-related compounds such as in E. chrysanthemi, where KdgT is involved in the uptake of 5-keto-4-deoxyuronate (DKI) and 2,5-diketo-3-deoxy-d-gluconate (DKII) (26) (Fig.1).

Biochemical Characterization of KDG Catabolism

Thorough kinetic investigations on whole cells and isolated membrane vesicles of E. coli K-12 demonstrated that the KdgT transporter is able to transport KDG (K _m = 0.25 mM) selectively and, to a lesser extent, d-glucuronate (K_m = 1.25 mM) (71, 72, 73). Both substrates can be concentrated several hundredfold inside the cells. KDG uptake is inhibited by classical electron transfer inhibitors, by proton-conducting compounds, and by thiol reagents (72). Uptake of the anionic sugars KDG and d-glucuronate involves the concomitant transport of protons (70) and is driven by the two components, ΔpH and ΔΨ, of the proton motive force (69). Mutants affected in the kdgT gene, selected for conditional growth at acidic pH (6.0), exhibited the same proton:sugar stoichiometry and ΔΨ dependence of the transport process as the wild-type strain, but were severely altered in their kinetic parameters for the substrates at alkaline pH (77). KdgT is the prototype of the KDGT family of KDG transporters (TC #2.A.10) (Table 2).

A unique KDG kinase has been partially purified from d-glucuronate- and d-galacturonate-grown cells (111). KdgK has an optimal activity around pH 6, which is enhanced in the presence of the divalent cations Mn²⁺, Mg²⁺, and Co²⁺. Apparent K_ms for ATP and KDG are 1 mM each (32, 111). KdgK is highly specific for KDG, since several structurally related compounds do not show any inhibition of its activity.

In contrast to the other enzymes, the KDPG aldolase (KdgA or Eda) is not restricted to the hexuronide/hexuronate pathway. It is the second enzyme of the Entner-Doudoroff pathway and catalyzes an aldol cleavage of KDPG to form pyruvate and 3-phosphoglyceraldehyde (27, 38). The purified enzyme has a pH optimum at 7.6 and does not need a cofactor for its activity (112). It displays high specificity for KDPG, with a K_m of 0.2 mM. The substrate of the reverse reaction, 3-phosphoglyceraldehyde, and 6-phosphogluconate competitively inhibit the KDPG aldolase, with K_is of 0.4 mM and 0.8 mM, respectively. KDPG aldolase is a multifunctional enzyme and is also able to catalyze the interconversion of 2-keto-4-hydroxyglutarate (KHG) with pyruvate and glyoxylate (143). This activity was originaly attributed to a distinct enzyme. The nucleotide sequence of the corresponding gene was reported (95) and subsequently shown to be identical to that of the E. coli eda (kdgA) gene (37). KHG/KDPG aldolase might also participate in regulation of the intracellular level of glyoxylate, which is known to be an inhibitor of citric acid cycle enzymes, via β-decarboxylation of oxaloacetate (45). The KHG/KDPG aldolase is a trimer of 22-kDa subunits (143). The substrate tolerance displayed by KDPG aldolase is useful in catalysis of the stereospecific aldol addition of pyruvate to a range of unnatural electrophilic substrates. In an attempt to elucidate its reaction mechanism, a recent crystallographic study of KDPG aldolase was undertaken (1). Determination of the structure of the enzyme at a resolution of 1.95 Å with a bound intermediate allowed identification of the active site. Directed evolution of KDPG aldolase has been employed to engineer enzymes with improved catalytic efficiency, altered substrate specificity, and stereoselectivity (40, 146).

Organization and Regulation of Genes Involved in KDG Catabolism

The structural genes of enzymes of KDG metabolism are scattered over the E. coli chromosome (Fig. 2). Strains mutated in the kdgT and kdgK genes were isolated by spontaneous resistance to the KDPG toxicity of an eda mutant (109, 115). Mutations in the eda gene were obtained by chemical mutagenesis (108). Isolation of thermosensitive mutations affecting KDG uptake (74) or KDPG aldolase activity (114) allowed the unambiguous identification of the structural genes. Plasmid-borne kdgT restores KDG and d-glucuronate uptake in kdgT mutants, and it directs the synthesis of a specific 28-kDa polypeptide associated with the membrane fraction, confirming the direct role of KdgT in transport (79). The kdgT gene is located at min 88.4 (109), the kdgK gene is at min 79.3 (115), and the eda gene is at min 41.6 (108, 114). The eda gene is tightly linked to the other gene of the Entner-Doudoroff pathway, edd (Table 1 and Fig. 2).

Synthesis of KdgT, KdgK, and Eda is induced in the presence of galacturonate, glucuronate, tagaturonate, or fructuronate. However, these enzymes are not synthesized coordinately depending on the inducer, in agreement with the scattering of the corresponding genes on the chromosome (113). For example, d-gluconate is a good inducer of Eda synthesis, but it represses KdgK synthesis. Moreover, Eda is less sensitive to catabolite repression than is KdgK. Use of appropriate mutants demonstrated that KDG is the true inducer formed by the sequential conversion from its precursors. However, KDG, used as an exogenous substrate, cannot induce its own transport system KdgT in the wild-type strain and behaves like a noninducing substrate (113). A class of mutants (including thermosensitive ones) which allowed growth of E. coli on KDG was found simultaneously to derepress KdgT, KdgK, and Eda activities. The corresponding mutations are located in the regulatory gene kdgR at min 41.1 in the vicinity of eda (115). KdgR is a transcriptional repressor that belongs to the IclR family (Table 3). It is assumed to have different affinities for the operators of the three operons, in the increasing order eda, kdgK, and kdgT. It is interesting to note that sensitivity of these operons to catabolite repression increases in the same order (71, 113, 115). Comparative genomic approaches revealed the presence of binding sites for KdgR in the regulatory regions of kdgT and kdgK, but not in that of eda, and identified the E. coli kduID operon as belonging to the kdgR regulon. Two well-conserved CRP consensus recognition sequences are also present upstream of kdgT (132). However, the final characterization of this regulon must await a complete molecular analysis that demonstrates direct binding of the transcriptional factors to the operator regions. Such an analysis was performed with the E. chrysanthemi KdgR repressor. It allowed the determination of the KdgR DNA recognition sequence (50), which is apparently conserved in E. coli (132) (Table 3).

When E. coli is grown on d-gluconate, the edd and eda genes are cotranscribed from a promoter located upstream of edd, which has been proposed to be controlled by the GntR repressor (37, 96). The basal level of eda expression is high, regardless of the carbon source (41, 113). This high transcription is presumably due to the two (or three) eda-specific promoters located in the coding sequence of edd (37). One of them was hypothesized to respond to induction by d-glucuronate (through its internal conversion into KDG) as mediated by the KdgR regulatory protein. However, no KdgR binding site is present in this region. The other promoter was activated by growth on d-gluconate (27, 37), but no GntR binding site is observed upstream of eda. Thus, the mechanisms responsible for regulation of eda transcription remain unclear. The large number of factors controlling expression of eda is consistent with the different physiological roles of Eda.

Occurrence of the KdgR Regulon in Other Bacteria

In contrast to its restricted role in E. coli, the KdgR regulator has a wide range of targets in the plant pathogenic bacterium E. chrysanthemi, which is able to degrade pectin (polygalacturonate) and to use the resulting oligomers as a carbon source for growth (for reviews, see references 50 and 124). The KdgR regulon contains about 20 operons encoding various types of pectinases as well as proteins necessary for secretion of pectinases, transporters of pectic oligomers (oligogalacturonates) and of KDG, and intracellular enzymes involved in the cleavage of pectic dimers and the catabolism of unsaturated monomers (KduI, KduD, and KdgK enzymes which degrade unsaturated monomers, are also produced by E. coli [Table 1]). In vitro analysis demonstrated that KdgR directly interacts with the promoter regions of the regulated genes and allowed the characterization of the KdgR binding site (86). KDG is the main inducing molecule which interacts with KdgR, causing dissociation of KdgR from its operators. In addition, two other intermediates of pectin catabolism, 5-keto-4-deoxyuronate (DKI) and 2,5-diketo-3-deoxygluconate (DKII), are also able to act as inducers. The KdgR protein is conserved in Enterobacteriaceae and it is functionally interchangeable between species (62, 139). The availability of novel complete bacterial genomes, including that of E. chrysanthemi (http://www.tigr.org/tdb/mdb/mdbinprogress.html), has provided the opportunity to perform comparative genomic analyses designed to predict the existence of new KdgR-regulated genes. Several have been identified and their proposed regulation has been verified experimentally (131, 132). Interestingly, the role of KdgR is not restricted to the already known negative control of pectin catabolism, but also extends to the positive control of genes involved in gluconeogenesis and of another repressor gene, pecT, controlling pectate lyase synthesis.

Salmonella enterica serovar Typhi and Y. pestis possess the three genes for ketogluconate catabolism, kdgK, kduI, and kduD. Klebsiella pneumoniae has only kdgK and kduD, while kduI is absent (132). The kdgT gene, encoding the transporter of KDG, is absent in all three of these species. The organization of genes involved in pectin catabolism in Erwinia carotovora closely resembles that found in E. chrysanthemi (131). Y. pestis and Yersinia enterocolitica also possess some pectinase genes and genes involved in the transport and degradation of oligogalacturonates. Based on the composition of their respective KdgR regulons, Y. pestis and Y. enterocolitica should be able to degrade pectic oligomers, K. pneumoniae to degrade short oligomers, limited to dimers or trimers, and S. enterica serovar Typhi and E. coli K-12 to degrade the monomers DKI, DKII, and KDG. Notably, another recently sequenced E. coli strain, CFT073, has acquired a cluster of genes allowing it to use short oligomers (131).

HEXONATES

d-Gluconate

Catabolism of d-gluconate in E. coli occurs in four steps (Fig.1). d-Gluconate is first transported into the cell by one of the four gluconate transporters that exist in the bacteria (GntT, GntU, GntP, and IdnT) (Table 2) and is phosphorylated by one of the two kinases (GntK and IdnK) to form 6-phospho-d-gluconate (Table 1). This compound is then catabolized by the two enzymes of the Entner-Doudoroff pathway, the 6-phosphogluconate dehydratase Edd and the 2-keto-3-deoxy-6-phophogluconate aldolase Eda. The synthesis of the gluconate kinase GntK and of only two of the permeases (GntT and GntU) is inducible by d-gluconate. They form the GntI system or main system for gluconate metabolism (99, 140).

The two transporters GntT and GntU belong to the same family (GntP) of gluconate:H⁺ symporters (TC #2.A.8) (Table 2) (98). GntT is a high-affinity transporter (K_m = 6 μM), whereas GntU has low affinity for d-gluconate (K_m = 212 μM), but synthesis of both is induced at a very low concentration of d-gluconate (2 to 10 μM) (99, 140). Expression of both transporters is subject to cAMP-dependent catabolite repression. Expression of GntT shows a pronounced peak very early in the logarithmic phase. GntT^– mutants grown on gluconate have a prolonged lag phase. GntT could be important for growth at low concentrations of d-gluconate or when cells enter the early exponential phase, whereas GntU could be important during the mid to late exponential growth phase (99). A third gluconate transporter, GntP, has been characterized in E. coli. It can transport d-gluconate with a high affinity (K_m = 25 μM), but its synthesis is not induced by this compound (68). Transport of d-gluconate by GntP is inhibited by a wide range of substrates, (including d-glucuronate, maltose, lactose, sucrose, fructose, and galactose), suggesting that it has a fairly broad specificity. The gntP gene is transcribed divergently from the uxuAB operon, which encodes enzymes involved in glucuronate metabolism. Moreover, an UxuR binding site is present in the regulatory region of gntP (132) (Fig. 2), suggesting that GntP is more probably a transporter for hexuronates or hexonates rather than a specific gluconate transporter. GntK is an ATP-dependent thermoresistant kinase that functions as a dimer (59).

The genes for gluconate metabolism are grouped in three loci on the E. coli chromosome. The gntK and gntU operon at min 77.1 is immediately downstream of the regulatory gene gntR (140) (Fig. 2). Transcription of all three genes can occur from a constitutive promoter located upstream of gntR, but another promoter exists in front of gntK. This promoter is inducible by d-gluconate through GntR and activated by cAMP-CRP (60). In addition, transcriptional attenuation was observed after gntK, which results in down-regulation of synthesis of the low-affinity permease GntU according to the availability of d-gluconate (60). The gntT gene is localized at min 76.4 (61, 99). The edd and eda genes are found at min 41.6, downstream of zwf, which encodes glucose-6-phosphate dehydrogenase, an enzyme of the pentose phosphate pathway. A transcription terminator separates zwf and edd. The genes edd and eda are cotranscribed in a polycistronic mRNA from a gluconate-inducible promoter, but other promoters present in the edd coding region could allow exclusive expression of eda (37). Induction of the genes of the gluconate catabolic pathway (gntK, gntU, gntT, edd, and eda) by d-gluconate occurs through the GntR regulatory protein (96). GntR belongs to the LacI family of transcriptional regulators (Table 3). Binding of GntR to the regulatory regions of gntT and gntK has been demonstrated by gel shift assays (96, 141). Binding of GntR to the operators is suppressed by d-gluconate and also by 6-phospho-d-gluconate, at a 10-fold higher concentration (96). A consensus for the GntR binding site has been determined by comparison of the regulatory regions of GntR-regulated genes. This sequence, ATGTTA(N₄; G-C rich)TAACAT, is present twice in the regulatory regions of gntK, gntT, and edd (99) (Table 3). A mutation in this sequence prevents GntR binding (141).

l-Idonate

In the absence of a functional GntI system, d-gluconate can be metabolized by the GntII (subsidiary) system, which includes a fourth gluconate permease, IdnT (alternate name, GntW), and the thermosensitive gluconate kinase IdnK (alternate name, GntV) (4). The two genes idnT and idnK are in a cluster of genes that encodes two dehydrogenase-like enzymes and that was shown to form the idonate catabolic pathway (8). E. coli grows well in minimal medium containing l-idonate (doubling time, 1.4 h). l-Idonate enters the bacteria by the IdnT permease and is oxidized to 5-keto-gluconate by IdnD (Fig. 1). 5-Ketogluconate is then reduced by IdnO to d-gluconate, which is phosphorylated by IdnK. IdnT transports d-gluconate with high affinity (K_m, 60 μM) (98) although l-idonate is the physiological substrate (8). The idn genes are grouped at min 96.8 on the chromosome (Fig. 2). The idnK gene is transcribed divergently from an operon comprising idnD, idnO, idnT, and idnR (alternate name, gntH); this last encodes the regulator of the idn genes (9) (Table 1). Expression of these two transcriptional units is inducible by l-idonate or 5-ketogluconate, but an internal promoter allows a constitutive expression of idnR (141). IdnR is an activator of idn gene expression. l-Idonate or 5-ketogluconate has been proposed as a coactivator (8, 9). IdnR belongs to the LacI family of transcriptional regulators and has 53% identity with GntR (Table 3). The high degree of amino acid similarity between the two regulators allows reciprocal cross-regulation between the two catabolic pathways, i.e., a repression of the genes of the GntI pathway by IdnR in the presence of l-idonate and the control of the genes of the GntII pathway by GntR. The two regulators probably bind to the same regulatory elements (141, 142) (Fig. 2).

Ketogluconates

Although E. coli grows poorly in minimal medium containing 2-keto-d-gluconate, 2,5-diketo-d-gluconate, or 2-keto-l-gulonate as the sole carbon source, it synthesizes enzymes that allow their catabolism. The catabolism of these ketogluconates is connected to the l-idonate and d-gluconate pathways (Fig. 1). The product of the gene ghrB (trkA, yiaE), located at min 80.1, is a 2-ketoaldonate reductase (Table 1). It catalyzes the NADPH-dependent reduction of 2-keto-l-gulonate, 2,5-diketo-d-gluconate, and 2-keto-d-gluconate to l-idonate, 5-keto-d-gluconate, and d-gluconate, respectively (147) (Fig. 1). The E. coli ghrB gene is expressed constitutively (147). The products of the genes dkgA (yqhE) and dkgB (yafB) have a 2,5-diketo-d-gluconate reductase activity and catalyze the reduction of 2,5-diketo-d-gluconate to 2-keto-l-gulonate (148) (Fig. 1). Nothing is known about the regulation of dkgA and dkgB.

In E. chrysanthemi, pectin catabolism leads to the formation of 5-keto-4-deoxyuronate, which is isomerized to 2,5-diketo-3-deoxygluconate and then is reduced to 2-keto-3-deoxygluconate by the isomerase KduI and the oxidoreductase KduD (50). Although E. coli is not able to grow on pectin or oligogalacturonates as the sole carbon source, the genes kduI and kduD are present (Table 1) (Fig. 1). They could be involved in the catabolism of 5-keto-4-deoxyuronate or in a secondary pathway for KDG catabolism.

d-Galactonate

d-Galactonate is transported by the DgoT permease (a member of the anion:cation symporter family, TC #2.A.1.14.7) (Table 2) and is dehydrated to 2-keto-3-deoxy-d-galactonate by the dehydratase DgoD. This compound is phosphorylated by the product of the DgoK kinase, and the resulting 2-keto-3-deoxy-6-phospho-d-galactonate is cleaved by the DgoA aldolase to give 3-phosphoglyceraldehyde and pyruvate (29, 33) (Fig. 1). The genes encoding enzymes of galactonate metabolism are clustered at min 83.5 in an operon formed by dgoR, encoding the repressor of the GntR family which controls the expression of this pathway (Table 3), and dgoK, dgoA, dgoD, and dgoT (Fig. 2). d-Galactonate itself is the true inducer.

l-Galactonate and l-Gulonate

E. coli strains K-12 and W grow readily on l-galactonate as the sole carbon source, whereas E. coli C grows very poorly and E. coli B fails to grow. Analysis of mutants of the galacturonate pathway showed that the products of uxaB, uxaA, kdgK, and eda, but not uxaC, are required for the catabolism of l-galactonate. l-Galactonate is converted to d-tagaturonate in one step by an oxidoreductase that utilizes NAD⁺ as cofactor. Synthesis of this enzyme is induced by l-galactonate (Fig. 1) (30). l-Galactonate is able to induce the synthesis of the enzymes of the galacturonate pathway, even in a strain mutated for the l-galactonate oxidoreductase, indicating that l-galactonate is an inducer of ExuR, the repressor of galacturonate catabolism. In contrast, d-tagaturonate cannot act reciprocally as an inducer of l-galactonate oxidoreductase synthesis. Thus, l-galactonate induces the synthesis of all the enzymes necessary for its catabolism by interacting with two regulators: the uncharacterized regulator of the synthesis of the l-galactonate oxidoreductase and ExuR (30).

While E. coli W can grow overnight on l-gulonate as sole carbon source, the E. coli strains K-12, B, and C cannot. However, after the plates are incubated for 3 to 4 days, strain K-12 yields colonies that have gained the ability to grow on l-gulonate. Analysis of mutants of the glucuronate pathway showed that the products of uxuB, uxuA, kdgK, and eda, but not uxaC, are required for the catabolism of l-gulonate. l-Gulonate is converted to d-fructuronate in one step by an oxidoreductase that utilizes NAD⁺ as cofactor. Synthesis of this enzyme is induced by l-gulonate (Fig. 1) (31). The l-galactonate oxidoreductase is also able to transform l-gulonate into d-fructuronate. but its synthesis is not induced by l-gulonate. The genes encoding the l-galactonate oxidoreductase and the l-gulonate oxidoreductase have not been characterized.

Salmonella typhimurium.

All the genes involved in the catabolism of d-gluconate, l-idonate, d-galactonate, and ketogluconates described in E. coli exist in S. enterica serovar Typhimurium. Analysis of the serovar Typhimurium genome shows that their genetic organization is conserved. However, no biological characterization of these pathways has been described. Serovar Typhimurium is able to grow on l-gulonate but not on l-galactonate as the sole carbon source.

ALDARATES

E. coli is able to use the diacid sugars d-glucarate and galactarate (an achiral compound) as sole carbon source for growth. Early biochemical studies by Blumenthal and Fish (21) reported the enzymatic steps of the d-glucarate pathway. d-Glucarate and galactarate are dehydrated to 5-keto-4-deoxy-d-glucarate, which is cleaved by an aldolase into tartronate semialdehyde and pyruvate. Tartronate semialdehyde is reduced to d-glycerate, which is then phosphorylated (Fig. 1). While early studies (21) identified 3-phosphoglycerate as the end product, recent nuclear magnetic resonance studies suggested that 2-phosphoglycerate rather than 3-phosphoglycerate is formed (48). Thus, pyruvate and 2-phosphoglycerate are the final products of the d-glucarate/galactarate catabolism.

The first step of aldarate catabolism is catalyzed by two enzymes, which are specific for each substrate (20, 23). The d-glucarate dehydratase GudD catalyzes the dehydration of d-glucarate into 5-keto-4-deoxy-d-glucarate. Galactarate is metabolized by a different dehydratase, GarD. The last three steps of the aldarate pathway are common to both d-glucarate and galactarate catabolism (Table 1). They are catalyzed by the 5-keto-4-deoxy-d-glucarate aldolase GarL, the tartronate semialdehyde reductase GarR, and the d-glycerate kinase GarK.

Genes Involved in Aldarate Catabolism

The enzymes involved in d-glucarate and galactarate catabolism were identified only after a genomic analysis involving comparison of proteins of unknown function encoded by the E. coli genome with known enzymes and with enzyme families (48). The sequence of a d-glucarate dehydratase previously characterized in Pseudomonas putida was used to search for related proteins encoded by the E. coli chromosome (48). One homologue (80% identity), encoded by the gene ygcX, indeed catalyzes the efficient dehydration of d-glucarate to 5-keto-4-deoxy-d-glucarate. Purification of the d-glucarate dehydratase from E. coli grown on d-glucarate as the sole carbon source and determination of its N-terminal sequence confirmed that the d-glucarate dehydratase effectively corresponded to the protein encoded by ygcX, now called gudD. In order to search for the E. coli gene encoding 5-keto-4-deoxy-d-glucarate aldolase, two candidates encoding homologues of diacid sugar aldolases were identified (48). The 5-keto-4-deoxy-d-glucarate aldolase was also purified from E. coli cells grown on d-glucarate, and its N-terminal sequence was determined. The N-terminal sequence of one homologue of diacid sugar aldolases corresponded to that of the purified aldolase. The corresponding gene, yhaF, is now called garL.

The gudD and garL genes are located in two regions on the E. coli chromosome proposed to contain all the genes involved in d-glucarate and galactarate catabolism (Fig. 2). At min 62.9, the first region contains a potential operon of three genes, gudD (ygcX), preceded by a gene of unknown function named gudX (ygcY) which encodes a GudD homologue (72% identity), itself preceded by the gene gudP encoding a permease of the MFS superfamily (TC #2.A.2). GudP is responsible for the uptake of both d-glucarate and galactarate (Table 2). The second region, at min 70.5, contains the gene garD (yhaG), encoding a dehydratase, and a divergent operon which includes four genes, garP (yhaU, encoding another permease of the MFS superfamily [54% identity with GudP]), garL (yhaF, encoding the identified aldolase), and garR (yhaE) and garK (yhaD), which encode the last two steps of the pathway responsible for tartronate semialdehyde reduction and phosphorylation.

Biochemical Characterization of Aldarate Catabolism

The transport of d-glucarate in E. coli cells was shown to have a K_m of 16 μM and to be competitively inhibited by galactarate (130). However, this analysis was performed on wild-type cells which contain both GudP and GarP transporters. The specificity of these two transport systems was not analyzed. The presence of gudP in the operon including gudD first suggested that the GudP protein is involved in d-glucarate uptake. Similarly, the location of garP at the vicinity of garD suggested that the GarP protein is responsible for galactarate uptake. The fact that galactarate is still an inducer in a garP mutant suggested the existence of a second galactarate transporter, probably GudP. Moreover, we noticed that several enterobacteria such as S. enterica serovar Typhimurium and E. chrysanthemi contain all the genes involved in d-glucarate/galactarate catabolism except garP, suggesting that the presence of garP is not essential.

While galactarate is not a substrate of GudD, this enzyme can use either d-glucarate, or 5-keto-4-deoxy-d-glucarate, or l-idarate (a compound observed only in this catabolic pathway) as a substrate (48). Thus, GudD catalyzes dehydration of both d-glucarate and l-idarate as well as their interconversion via epimerization. The relative rates of these reactions were determined by nuclear magnetic resonance (48), which showed that the rate of dehydration slightly exceeds that of epimerization. GudD has an efficient activity with d-glucarate or l-idarate as a substrate, with K_ms of 60 and 16 μM, respectively. These two compounds are poor substrates for the dehydratase GudX, encoded by the gene adjacent to gudD. Since other substrates could not be identified, the physiological role of GudX remains unclear. GudX could be the result of a gene duplication leading to new substrate specificity towards an unidentified sugar diacid. We noticed that the gudX gene is well conserved, adjacent to gudD, in other enterobacteria, such as S. enterica serovar Typhimurium and E. chrysanthemi. This conservation suggests that GudX indeed has a novel functionality. GudD and GudX are members of the mandelate racemase subgroup of the enolase superfamily. The dehydration reactions as well as the epimerization of d-glucarate and l-idarate can be rationalized by the participation of a common enolic intermediate. Crystal structures of E. coli GudD have been obtained (43, 44). GudD is a tetramer of four identical monomers containing two domains. An N-terminal domain that consists of three β-sheets and three α-helices precedes a C-terminal β/α-barrel domain. The structure of GudD reveals the three residues involved in Mg²⁺ binding and the presence of the catalytic bases involved in proton abstraction. 4-Deoxy-d-glucarate is a competitive inhibitor of GudD activity (43, 44).

Purified galactarate dehydratase was shown to be unstable (22, 23). Hubbard et al. (48) showed that GarD could be reactivated and assayed using conditions similar to those described for the homologous dehydratase UxaA (32% identity) involved in galacturonate catabolism. GarD has maximal activity in the presence of 0.5 mM Fe^{2 +} and 170 mM β-mercaptoethanol at pH 7.5. The K_m of GarD for galactarate is 0.8 mM.

The functions of GarL, GarR, and GarK were characterized after purification of the recombinant proteins with an N-terminal His tag (48). Successive enzymatic reactions using 5-keto-4-deoxy-d-glucarate as a substrate provide evidence that these three proteins are involved in the three last steps of the d-glucarate/galactarate catabolic pathway. The K_ms of the aldolase GarL and the kinase GarK for their substrates are 65 μM and 51 μM, respectively (48). The instability of the oxidoreductase GarR means it is difficult to determine its kinetic constants. GarL belongs to the class II aldolases, which use divalent metal ions as an electron sink for the cleavage/addition reaction. Indeed, GarL requires a divalent cation (such as Mg²⁺, Co²⁺,or Mn²⁺) for catalysis (39). In organic chemistry, aldolases are useful catalysts for the stereo-controlled formation of carbon-carbon bonds. As with Eda, GarL has raised some interest due to its low substrate specificity and its ability to condense a wide range of aldehydes with pyruvate. The crystal structure of GarL was determined in an attempt to provide information on its reaction mechanism (10, 58). In solution, the enzyme functions as a hexamer (subunit molecularmass of 27.4 kDa). The structure of the monomer adopts a modified (α/β)₈ barrel fold. The crystal structure revealed two phosphate anionsbound within the active site pocket, suggesting thatphosphate may play a role in the enzymatic activity and/or cation binding.

Regulation of Aldarate Catabolism

Early studies revealed that growth on d-glucarate induced both d-glucarate and galactarate catabolism (21). Recently, Monterrubio et al. (85) showed that the synthesis of the enzymes GudD, GarD, GarK, and GarR is induced by growth on galactarate, d-glucarate, or their common intermediate, d-glycerate. In each case, d-glycerate was the better inducer. Construction of lacZ fusions indicated that functional promoters are located 5' of the genes garD, garP, and gudP (Fig. 2). Polarity effects of insertion mutations confirmed that genes garPLRK are cotranscribed. Use of gene fusions in each operon confirmed that their expression is induced in the presence of galactarate, d-glucarate, or d-glycerate. d-Glycerate is two- to threefold more efficient than the two other substrates (85).

Isolation of a mutation affecting growth on d-glucarate and galactarate, but located outside the gar and gud loci, led to the identification of a gene involved in the regulation of pathway synthesis (85). The mutation was situated at min 3.9, in the gene yaeG, first renamed sdaR and now called cdaR (Fig. 2). Use of gene fusions showed that the expression of the cdaR gene is positively autoregulated. In the presence of inducers, CdaR is activated, thereby increasing its own expression and that of the three structural gene operons garD, garPLRK, and gudPXD. The amino acid sequence of CdaR is not significantly similar to that of previously reported regulatory proteins (Table 3). CdaR could be the prototype for a new class of transcriptional regulators, but the direct interaction between CdaR and DNA remains to be demonstrated.

Acknowledgments

Studies in this laboratory were performed at the Institut National des Sciences Appliquées de Lyon and were mainly supported by grants from the Centre National de la Recherche Scientifique and the Ministère de l'Education Nationale et de la Recherche.