INTRODUCTION

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Next to glucosyl residues, the benzene ring is the most widely distributed unit of chemical structure in nature (59, 132). The complex aromatic polymer lignin comprises about 25% of the land-based biomass on Earth, and the recycling of this and other plant-derived aromatic compounds is vital for maintaining the Earth's carbon cycle. The degradation of such chemicals is accomplished mainly by microorganisms (129, 321), and in recent years there has been considerable interest in exploring their ability to degrade and detoxify the increasing amounts of aromatic compounds which enter the environment as by-products of many industrial processes (239). Although many genera of microorganisms degrade aromatic compounds other than aromatic amino acids, with Pseudomonas being the most extensively analyzed (132, 335), this ability has been only occasionally studied in enteric bacteria. The early literature contains reports on the formation of phenol and p-cresol by Escherichia coli bacterial cultures growing in natural media, peptone and casein media, and in chemically defined media containing L-tyrosine or p-hydroxybenzoic acid (316). The decarboxylation of substituted cinnamic acids with the production of volatile phenolic compounds that provide phenolic flavors in fermented beverages has been reported for many enterobacteria of the genera Klebsiella, Enterobacter, and Hafnia, as well as for Escherichia intermedia (179). The use of benzoic acid, p-hydroxybenzoic acid, and phenylacetic acid (PA) by Enterobacter aerogenes as the sole carbon and energy source was one of the first reports of the catabolism of aromatic acids by an enterobacterium (119). In the mid-1970s, Chapman and coworkers found that most enterics could use aromatic compounds, with the ability to utilize hydroxyphenylacetic acid (HPA) being the most widespread (38). It was the work of Cooper and Skinner in 1980 on the ability of E. coli to mineralize 3- and 4-hydroxyphenylacetic acids (3HPA and 4HPA) that delineated for the first time a complete catabolic pathway in enteric bacteria (50). Later, Burlingame and Chapman (36) reported that many laboratory strains and clinical isolates of E. coli can catabolize various aromatic acids. A historical perspective on the mineralization of aromatic compounds by E. coli is summarized in Table 1.

At first view, the ability of E. coli to degrade aromatic compounds appears to be an unexpected finding since this feature has always been associated with typical soil bacteria and E. coli is mainly regarded as an inhabitant of the animal gut. However, when one analyzes the ecology of E. coli, it becomes clear that this bacterium may easily encounter aromatic compounds in both the intestinal and extraintestinal habitats that it colonizes, which explains its catabolic potential to use such compounds as carbon and energy source.

The intestine of warm-blooded animals, the primary habitat of E. coli, contains some 400 to 500 different bacterial species, with E. coli being the most abundant (making up about 1% of the total fecal bacterial flora) (233). The success of E. coli in the gut ecosystem is thought to reflect its abilities to occupy different ecological niches. Thus, since E. coli grows both anaerobically and aerobically, it is able to colonize intestinal habitats in which oxygen offers some ecological advantage. Such habitats could be ones in close proximity to epithelial cells, where oxygen molecules might pass from the blood through the epithelium to the microbes attached to it. By assimilating such molecules, E. coli may be important in developing and maintaining the oxygen-free conditions and low oxidation-reduction potential favoring strict anaerobes in the large intestine (280). While embedded within the mucus layer overlying intestinal epithelial cells, E. coli grows with a generation time of 40 to 80 min (245, 246). In contrast, the population of E. coli cells in the cecal luminal contents are essentially static with respect to growth and are excreted in the feces (246).

Although E. coli is a highly successful commensal of the intestines of warm-blooded animals as well as a pathogen of the enteric, urinary, pulmonary, and nervous systems, this facultative anaerobe must survive and grow outside the animal host to effect successful interhost spread. E. coli is not famous for extracorporeal existence, but it nonetheless shares with its soil-inhabiting relatives such as Klebsiella species the ability to thrive under a wide range of various physical and chemical conditions, including adaptation for survival over long periods of nongrowth (210). Soil, water, sediment, and perhaps food are other habitats of E. coli, and the bacterium might spend comparable times in each of its two main habitats (88, 281). While pollution from human sources may be the most important source of E. coli in the environment (29, 260), the fact that this bacterium was found in pristine tropical waters, where it remained physiologically active and grew at rates dependent on nutrient levels, suggests that it can be a natural inhabitant in these environments and that it may be part of a previously established community (22). E. coli can also replicate and survive in soil protozoa. Since protozoa are widely distributed in soils and effluents, they may also constitute an environmental reservoir for transmission of this enterobacterium (17). Thus, unlike host-specific or obligate parasites, E. coli is a highly adaptable microorganism with an extensive repertoire of metabolic and regulatory genes that facilitate the colonization of widely different environments (78).

As stated above, aromatic compounds are highly abundant in soil and water, and therefore it is obvious that they can constitute a normal carbon source for E. coli when this bacterium reaches its extraintestinal habitat. Although it is still not known which substrates E. coli grows on in the large intestine and which pathways provide it with the metabolic advantage necessary for it to compete with the hundreds of other bacteria with which it shares this habitat, it is likely that aromatic compounds can also be a frequent carbon source for E. coli in the animal gut. Aromatic amino acids and plant constituents are the major sources of aromatic compounds in the gastrointestinal tract. Minor sources of aromatic compounds in the human gut include certain steroids and some drugs and food constituents (additives, colorants, and contaminants) (117). Estimates suggest that between 3 and 25 g of protein and peptides enters the large bowel every day from the diet, as well as from endogenous sources such as host tissues, bacterial debris, pancreatic enzymes, and other secretions (296). In the large gut, these substances are depolymerized by a mixture of residual pancreatic endopeptidases and bacterial proteases and peptidases. The resulting short peptides and amino acids then become available for fermentation by many intestinal anaerobes such as Bacteroides, Lactobacillus, Bifidobacterium, Clostridium, and Streptococcus species, generating a wide range of phenolic and indolic compounds in a series of deamination, transamination, decarboxylation and dehydrogenation reactions. Phenol, p-cresol, HPA, hydroxyphenylpropionic acid (HPP), and hydroxybenzoic acid are the principal products of tyrosine fermentation in the human large intestine, while PA, phenylpropionic acid (PP), and benzoic acid are produced from phenylalanine. PP may, however, also be formed from tyrosine (296, 320). Phenylethylamine (PEA) and tyramine are produced from decarboxylation of phenyalanine and tyrosine, respectively (320). Since the production of these aromatic compounds was inhibited in the presence of a readily fermentable source of carbohydrate, carbohydrate availability may be a critical factor affecting aromatic amino acid fermentation in the large intestine (296). The aromatic compounds generated by the intestinal anaerobes meet a variety of fates in the body. Thus, they may be detoxified by glucuronide or sulfate conjugation or may remain unabsorbed and be voided in the feces and urine. They can also completely break down under local aerobic conditions in the large intestine as a result of the action of some facultative anaerobes such as E. coli, such that mono- and dioxygenases are able to incorporate molecular oxygen into the aromatic ring (296, 320). A second major source of aromatic compounds in the animal gut involves the dietary plant constituents such as ferulic and caffeic acid, which result in HPP acids (235), as well as different flavonoid glycosides that are ingested in daily quantities of 1 to 2 g by humans. A collaborative bacterial catabolism, i.e., syntrophic interactions among microorganisms, of flavonoids in the human gut has been suggested. Thus, a number of obligately anaerobic bacteria from the human intestinal flora, e.g., Bacteroides species, are capable of cleaving the glycosidic bond of flavonoids, generating the corresponding aglycones such as quercetin, kaempferol, naringenin, and catechin (338). These aglycones are then subject to ring cleavage by different bacteria, e.g., Clostridium and Eubacterium species, giving rise to a variety of aromatic acids, such as 4HPA (from kaempferol), 3,4-dihydroxyphenylacetic acid (from quercetin), PA (from naringenin), and HPP (from tricetin and tricin) (121, 285, 338), that can be utilized by other intestinal bacteria such as E. coli.

Phylogenetic analyses have shown that E. coli strains fall into four main phylogenetic groups (A, B1, B2, and D), where virulent extraintestinal strains belong mainly to groups B2 and D, whereas most commensal strains belong to group A (46). E. coli K-12 (group A) is by far the most extensively studied E. coli strain, and it represents the best-understood living organism at the biochemical and genetic levels. The complete sequences of the 4.6-Mb genome of two E. coli K-12 derivatives, MG1655 (27; EcoGene database accessible using the Colibri website, http://www.genolist.pasteur.fr/Colibri/) and W3110 (202; GenoBase database accessible at the website http://ecoli.aist-nara.ac.jp/), have been reported, and functional genomic analyses are being performed (291). The wild-type strain of E. coli K-12 was isolated from the feces of a convalescent diphtheria patient in 1922 at Stanford University, and subcultures and derivatives of this strain were first reported in 1944 (120). Since the K-12 strains are unable to colonize the human gut (297), the K-12 lineage is considered to be the prototype of a biologically safe vehicle for the propagation of many efficient gene-cloning and expression systems. However, the presence of K-12 strains among E. coli isolates is extremly low, since no K-12 strains were detected among 226 environmental and human stool isolated samples (29). Other E. coli laboratory strains not derived from K-12 are, for instance, E. coli C, B, and W. E. coli C (no. 122 of the National Collection of Type Cultures, London, United Kingdom) is a prototroph F strain (24) and was one of the first strains shown to be able to recombine (acting as a recipient) with K-12 (177). E. coli B is a wild-type E. coli strain (67). A mutant of strain B that is resistant to radiations, E. coli B/r, has been frequently used in laboratory studies (339). The W (or Waksman) wild-type strain of E. coli (ATCC 9637) was apparently isolated from the soil of a graveyard by S. A. Waksman (E. Ron, personal communication). A vitamin B₁₂ auxotroph derivative of the W wild-type strain, E. coli ATCC 11105 (hereafter referred as E. coli W) (63), is a well-known penicillin G acylase producer. During the writing of this review, the 5.4-Mb genome of the enterohemorrhagic E. coli O157:H7 strain (group D) was reported (236). The genomes of E. coli K-12 and O157:H7, two strains that last had a common ancestor about 4.5 million years ago, revealed an unexpectedly complex segmented relationship (236). This diversity within the E. coli species is reflected in significant differences in gene content among different strains whose chromosome size ranges from 4.5 to 5.5 Mb (214, 262). Genes found in most individuals, that is, the core set of genes for that species (core gene pool), are the genes that determine those properties characteristic of all members of the species. Additionally, each strain has auxiliary genes distributed throughout the genome (flexible gene pool), e.g., pathogenicity islands and metabolic pathway genes, which determine properties found in some but not all members of the species and that contribute to maintaining the dynamic gene pool in E. coli (78, 125, 171, 236, 262). In this sense, the E. coli intraspecies variation in the ability to use different aromatic acids as sole carbon and energy sources (36) (Table 2) is a clear example that E. coli strains possess different capacities for utilizing growth-limiting nutrients and that they are likely to be used to increase the fitness and to expand the ecological niches of individual E. coli cells. The work of Burlingame and Chapman (36) revealed that all seven laboratory strains tested grew using PP, 3HPP, or 3-hydroxycinnamic acid (3HCI) as the sole carbon and energy source. However, while E. coli W is also able to grow on PA and 4HPA or 3HPA, E. coli K-12 and E. coli NCTC 5928 grew on PA but not on 4HPA or 3HPA and E. coli B and E. coli C grew on 4HPA or 3HPA but not on PA (Table 2). None of the strains tested was able to grow on 2HPA, cinnamic acid (CI) or its 2- or 4-hydroxy derivatives, or with the 2- or 4-hydroxy derivatives of PP (36). Among the clinical isolates analyzed, 48 (72%) of 67 could grow on at least one of the six aromatic acids tested and 27 (40%) could use all six (Table 2). Strains that could grow on 4HPA could also grow on 3HPA, and those that grew on 3HPP also grew on 3HCI, consistent with a common catabolic pathway for each of these two pair of compounds (see below).

The aim of this article is to review the genetics and biochemistry of the catabolism of aromatic compounds in E. coli. Our understanding of the utilization of these compounds by E. coli has leapt forward in recent years with the genetic characterization of the cognate catabolic pathways. Additional information has become available through the sequencing of the E. coli genome. Homologues of many of the E. coli genes involved in the catabolism of aromatic compounds can be identified on other bacterial genomes, some of which have been completely sequenced at the time of writing. Where possible, data derived from these genomes have been also included in this article and some evolutionary considerations have been pointed out. Finally, the use of E. coli as a biocatalyst for biotransformation or biodegradation of aromatic compounds will be addressed. Conclusions derived from this review may provide useful starting points for future research.

As indicated in the Introduction, E. coli is able to mineralize several aromatic compounds (Table 2). In this section, the general organization of the gene clusters for the degradation of 4HPA, 3HPA, 3HPP, 3HCI, PP, PA, and some aromatic amines is revised.

Some E. coli laboratory strains such as E. coli B, C, and W, but not E. coli K-12, are able to degrade 4HPA, 3HPA, and homoprotocatechuate (3,4-dihydroxyphenylacetate [HPC]) via an inducible chromosomally encoded meta-cleavage pathway (36, 50). The genes encoding the HPC meta-cleavage degradative route from E. coli C (hpc cluster) and E. coli W (hpa cluster) have been cloned and sequenced (143, 250, 270). The upper hpa gene cluster encoding the enzymes responsible for the hydroxylation of 4HPA and 3HPA to the catecholic intermediate HPC in E. coli W has also been cloned and sequenced, and it is located immediately adjacent to the HPC meta-cleavage genes (Fig. 1A). The homologous upper hpa cluster of E. coli C has also been cloned and partially sequenced (251, 254). In addition, near the hpa cluster E. coli W contains the pac gene (Fig. 1A), which encodes penicillin G acylase, an enzyme able to hydrolyze a wide range of amides and esters of HPA and PA (251, 254) (see below). The biochemical pathway for the catabolism of HPC in E. coli W (250) is similar to that previously delineated in E. coli C (50, 143, 270) (Fig. 1B). The HPC dehydrogenative route in E. coli W involves meta-cleavage of HPC by HPC 2,3-dioxygenase (HpaD), to give 5-carboxymethyl-2-hydroxymuconic semialdehyde (CHMS), which undergoes dehydrogenation to 5-carboxymethyl-2-hydroxymuconic acid (CHM) by the action of the CHMS dehydrogenase (HpaE). CHM isomerizes to 5-oxo-pent-3-ene-1,2,5-tricarboxylic acid (OPET) through a CHM isomerase (HpaF), and the OPET undergoes decarboxylation to 2-hydroxy-hept-2,4-diene-1,7-dioic acid (HHDD) by the HpaG decarboxylase. The formation of 2,4-dihydroxy-hept-2-ene-1,7-dioic acid (HHED) through the action of the HpaH hydratase on the product of the HpaG-catalyzed reaction is followed by its HpaI-mediated cleavage to give pyruvate and succinic semialdehyde as final products (Fig. 1B). Although pyruvate is a central metabolite, succinic semialdehyde requires previous dehydrogenation to succinic acid to enter the Krebs cycle, and this enzymatic step is not encoded within the hpa cluster (see below).

View larger version (23K): [in this window] [in a new window]   FIG. 1.   Pathway for the catabolism of HPA (4HPA and 3HPA) in E. coli. (A) Genetic map of the chromosomal hpa (in E. coli W) and hpc (in E. coli C) regions. Relevant genes are indicated by blocks: genes with similar shading participate in the same enzymatic step or in the same functional unit (route) of the pathway. The hpc genes are indicated in brackets. Regulatory and transport genes are shown by solid and vertically striped blocks, respectively. The genes flanking the hpa cluster (tsr, orf12, orf13, and orf14) are contiguous in the genome of E. coli K-12 and are represented by thick lines. orf12 and orf13 correspond to the yjiY gene from E. coli K-12. orf14 corresponds to the yjiA gene from E. coli K-12. The arrows show the directions of gene transcription. Bent arrows represent the Pr, Pg, Px, Pa1, Pa2 and PBC promoters. REP sequences are shown. The HpaR repressor and HpaA activator are represented by a square and hexagon, respectively; empty and solid symbols indicate inactive and active regulators, respectively;  and + indicate transcriptional repression and activation, respectively. The inducer molecule (HPA and HPC) is represented by a solid circle. (B) Biochemistry of the HPA catabolic pathway. The metabolites are 4HPA and 3HPA, HPC (homoprotocatechuate), CHMS (5-carboxymethyl-2-hydroxy-muconic semialdehyde), CHM (5-carboxymethyl-2-hydroxy-muconic acid), OPET (5-oxo-pent-3-ene-1,2,5-tricarboxylic acid), HHDD (2-hydroxy-hept-2,4-diene-1,7-dioic acid), OHED (2-oxo-hept-3-ene-1,7-dioic acid), and HHED (2,4-dihydroxy-hept-2-ene-1,7-dioic acid). The enzymes are HpaBC (HPA monooxygenase), HpaD (HPC 2,3-dioxygenase), HpaE (CHMS dehydrogenase), HpaF (CHM isomerase), HpaG (OPET decarboxylase), HpaH (hydratase), HpaI (HHED aldolase), and Sad (succinic semialdehyde dehydrogenase). The HPA transport protein (HpaX) is represented by a thick arrow. Out and In indicate outside and inside the cell, respectively.

The hpa catabolic cluster of E. coli W (Fig. 1A) is composed of 11 genes arranged as follows: (i) eight enzyme-encoding genes organized in two putative operons, the 4HPA/3HPA hydroxylase operon (hpaBC) and the HPC meta-cleavage operon (hpaGEDFHI) homologous to the hpcECBDGH operon from E. coli C; (ii) two regulatory genes, hpaR (hpcR in E. coli C) and hpaA, that control the expression of the meta-cleavage and HPA hydroxylase operons, respectively; and (iii) the hpaX gene, encoding the 4HPA/3HPA transport protein. All the genes are transcribed in the same direction with the sole exception of hpaR (250) (Fig. 1A). Analysis of the intergenic regions revealed the presence of five REP (repetitive extragenic palindromic) or PU (palindromic unit) sequences (16). While two REP sequences are located in the largest intercistronic region of the meta-cleavage operon between the hpaF and hpaH genes, the other three REP sequences are found at the 3' end of such operon and might act as transcription termination signals (Fig. 1A). Two putative hairpin loops that might act as transcriptional terminators are also located downstream of the hpaA gene and hpaBC operon (250).

Analysis of the regions flanking the hpa cluster from E. coli W showed the presence of several open reading frames (orf genes) (250) (Fig. 1A). Downstream of hpaR is located an incomplete orf that corresponds to the tsr gene mapped in the chromosome at 98.9 min (hereafter, chromosomal mapping in E. coli refers to that of E. coli K-12 strain MG1655 [EcoGene database at the Colibri website]) and that encodes the serine chemoreceptor (6). At the other end of the hpa cluster, that is, downstream of the hpaC gene, there are several orf genes that correspond to the yjiY (orf12 and orf13) and yjiA (orf14) genes of E. coli K-12 (EcoGene database at the Colibri website) (Fig. 1A).

Most E. coli strains are able to degrade 3HPP via a meta-fission pathway (36) (Table 2), and several mutants defective in this catabolism have been isolated and characterized (39). The biochemical pathway for the catabolism of 3HPP in E. coli K-12 (36, 39) is shown in Fig. 2B. The first step is catalyzed by the MhpA hydroxylase, which inserts one atom of molecular oxygen into the 2 position of the phenyl ring of 3HPP to give 2,3-dihydroxyphenylpropionate (DHPP), which, when acted on by 2,3-dihydroxyphenylpropionate 1,2 dioxygenase (MhpB), undergoes an extradiol cleavage, yielding the meta ring fission product, 2-hydroxy-6-keto-nona-2,4-diene 1,9 dioic acid (HKNDA). HKNDA is cleaved by the MhpC hydrolase to give succinate and 2-hydroxy-penta-2,4-dienoic acid (HPDA), which is hydrated to 4-hydroxy-2-ketopentanoic acid (HKP) by the MhpD hydratase. Then the MhpE aldolase catalyzes the fission of HKP to give pyruvate and acetaldehyde, with the latter being converted to acetyl coenzyme A (acetyl-CoA) through the action of the MhpF acetaldehyde dehydrogenase (Fig. 2B).

View larger version (19K): [in this window] [in a new window]   FIG. 2.   Pathway for the catabolism of 3HPP in E. coli. (A) Genetic map of the chromosomal mhp cluster. Relevant genes are indicated by blocks: genes with similar shading participate in the same enzymatic step or in the same functional unit (route) of the pathway. Regulatory and transport genes are shown by solid and vertically striped blocks, respectively. Genes flanking the mhp cluster (lacI and yaiL) are represented by thick lines. The arrows show the directions of gene transcription. Bent arrows represent the Pr and Pa promoters. The location of the BIME is shown. The inactive and active forms of the MhpR activator are represented by empty and solid hexagons, respectively. + indicates transcriptional activation. The inducer molecule (3HPP and 3HCI) is represented by a solid circle. (B) Biochemistry of the 3HPP catabolic pathway. The metabolites are 3HPP, DHPP (2,3-dihydroxyphenlypropionic acid), HKNDA (2-hydroxy-6-keto-nona-2,4-diene 1,9-dioic acid), HPDA (2-hydroxy-penta-2,4-dienoic acid), and HKP (4-hydroxy-2-ketopentanoic acid). The enzymes are MhpA (3HPP monooxygenase), MhpB, (DHPP 1,2 dioxygenase), MhpC (HKNDA hydrolase), MhpD (HPDA hydratase), MhpE (HKP aldolase), and MhpF (acetaldehyde dehydrogenase [acylating]). The 3HPP transport protein (MhpT) is represented by a thick arrow. Out and In indicate outside and inside the cell, respectively.

Analysis of the 9.8-kb mhp cluster (92) (Fig. 2A) reveals the existence of eight genes arranged as follows: (i) six genes encoding the initial monocomponent monooxygenase (mhpA), the extradiol dioxygenase (mhpB), and the hydrolytic meta-cleavage enzymes (mhpCDFE); (ii) one regulatory gene (mhpR); and (iii) one gene (mhpT) that encodes a transporter and is flanked by two bacterial interspersed mosaic elements (BIMEs) that belong to the BIME-2 subfamily (16). All the genes appear to be transcribed in the same direction, with the sole exception of mhpR (Fig. 2A). Although the Shine-Dalgarno sequences of the mhpR, mhpA, and mhpC genes are located in intergenic regions, those of mhpB, mhpD, mhpF, and mhpE overlap the preceding genes, suggesting that the most common mechanism of translational coupling may occur (92).

The mhp cluster maps at min 8.0 of the chromosome, between the lacI gene, encoding the transcriptional repressor of the lac operon, and the adhC gene, encoding an alcohol-acetaldehyde dehydrogenase, that is in the vecinity of the tau operon for taurine metabolism (EcoGene database at the Colibri website). The order of the catabolic genes in the meta-cleavage operon, with the single exception of mhpF, parallels that of the enzymatic steps in the 3HPP catabolic pathway (92) (Fig. 2).

It is known that E. coli is also able to grow with 3HCI as the sole carbon and energy source (36) (Table 2). Growth with 3HCI induces the synthesis of the MhpA and MhpB enzymes, which are responsible for the initial attack on 3HPP (36), and it has been shown that the mhp cluster from E. coli confers to Salmonella enterica serovar Typhimurium LT2, a strain unable to grow on 3HPP and 3HCI, the ability to use these two aromatics as the sole carbon and energy source (92). Hence, the mhp genes are also responsible for the catabolism of 3HCI via 2,3-dihydroxycinnamic acid (DHCI) with the formation of pyruvate, acetyl-CoA, and fumarate (Fig. 3).

View larger version (13K): [in this window] [in a new window]   FIG. 3.   Biochemistry of the 3HCI catabolic pathway. The metabolites are 3HCI, DHCI (2,3-dihydroxycinnamic acid), HKNTA (2-hydroxy-6-keto-nona-2,4,7-triene 1,9-dioic acid), HPDA (2-hydroxy-penta-2,4-dienoic acid) and HKP (4-hydroxy-2-ketopentanoic acid). The enzymes are MhpA (3HCI monooxygenase), MhpB, (DHCI 1,2-dioxygenase), MhpC (HKNTA hydrolase), MhpD (HPDA hydratase), MhpE (HKP aldolase), and MhpF (acetaldehyde dehydrogenase [acylating]). The 3HCI transport protein (MhpT) is represented by a thick arrow. Out and In indicate outside and inside the cell, respectively.

The catabolism of PP in E. coli is initiated by a dioxygenolytic pathway (36, 39) (Fig. 4B). The first step is catalyzed by a PP dioxygenase, which inserts an atoms of molecular oxygen into each of positions 2 and 3 of the phenyl ring of PP, yielding cis-3-(3-carboxyethyl)-3,5-cyclohexadiene-1,2-diol (PP-dihydrodiol), which is subsequently oxidized by the PP-dihydrodiol dehydrogenase to give DHPP (36, 39) (Fig. 4B). DHPP is the substrate of the meta-cleavage pathway described above for 3HPP degradation, and therefore it links the catabolism of PP and 3HPP in E. coli. The nucleotide sequence of a 7.2-kb DNA fragment that carries the hca cluster for PP catabolism revealed (i) five genes encoding the PP-dioxygenase (hcaEFCD; formerly named as hcaA1A2CD) and PP-dihydrodiol dehydrogenase (hcaB), (ii) a regulatory gene (hcaR), and (iii) a gene (hcaT) that might encode a transporter (71). Downstream of hcaD, the closely linked orfX (Fig. 4A) could also belong to the hca cluster, and it codes for a 155-amino-acid (aa) product of unknown function. The Shine-Dalgarno sequences of hcaE, hcaF, hcaC, hcaB, and hcaD overlap the preceding genes, suggesting that translational coupling occurs (71). Immediately downstream of orfX there is an inverted repeat sequence that could act as a transcriptional terminator of a potential operon. hcaR and hcaT are located upstream of hcaEFCBDorfX, but they are transcribed in the opposite direction from the other genes (Fig. 4A). Although the intergenic spacing between hcaR and hcaT was 159 bp, no typical transcriptional terminator and promoter sequences were detected in this DNA fragment (71).

View larger version (16K): [in this window] [in a new window]   FIG. 4.   Pathway for the catabolism of PP in E. coli. (A) Genetic map of the chromosomal hca cluster. Relevant genes are indicated by blocks: genes with similar shading encode the subunits of the PP dioxygenase. Regulatory (solid block) and putative transport (vertically striped block) genes are also shown. The horizontally striped block indicates the gene encoding the PP dihydrodiol dehydrogenase. The empty block represents a gene of unknown function. The csiE gene flanking the hca cluster is represented by a thick line. The arrows show the directions of gene transcription. Bent arrows represent the Pr and Pe promoters. The inactive and active forms of the HcaR activator are represented by empty and solid hexagons, respectively. + indicates transcriptional activation. The inducer molecule (PP and CI) is represented by a solid circle. (B) Biochemistry of the PP catabolic pathway. The metabolites are PP, CI, PP dihydrodiol, CI-dihydrodiol, DHPP, and DHCI (see the legends to Fig. 2 and 3). The enzymes are HcaEFCD (PP dioxygenase), HcaB (PP-dihydrodiol dehydrogenase), and MhpBCDEF (see the legend to Fig. 2). The putative PP/CI transport protein (HcaT) is represented by a thick arrow. Out and In indicate outside and inside the cell, respectively.

The hca cluster maps immediately downstream of csiE, which encodes the stationary-phase-inducible protein CsiE, at 57.5 min of the E. coli chromosome (EcoGene database at the Colibri website) and therefore far from min 8, where the mhp cluster responsible for DHPP degradation is located (see above).

The HcaEFCD dioxygenase and HcaB dihydrodiol dehydrogenase are able to transform CI into DHCI (Fig. 4B) (36, 71). Thus, when the hca and mhp clusters from E. coli are expressed in S. enterica serovar Typhimurium LT2, the resulting strain can grow on minimal medium containing CI as the sole carbon and energy source (71). Therefore, while in some soil Pseudomonas species and in Lactobacillus pastorianus the catabolism of CI could be accomplished by an initial reduction of the double bond of the side chain with formation of PP, the catabolism of CI by the Hca enzymes in serovar Typhimurium produces DHCI, which, following the mhp-encoded pathway, will be finally mineralized to pyruvate, acetyl-CoA, and fumarate (71) (Fig. 3). Since CI can induce the expression of the hca genes and is converted to DHCI by E. coli cells in a resting-cell process, it is difficult to explain the unexpected observation that this bacterium does not grow when cultivated on CI as the sole carbon and energy source (71). A possible explanation of this lack of growth could be that the DHCI generated in the reactions catalyzed by the Hca enzymes (and not the DHCI generated from 3HCI by the MhpA hydroxylase) or other intermediates farther down the mhp-encoded pathway accumulate to a toxic level that prevents the normal metabolic flux of the cell (71). Why this toxicity is not observed in serovar Typhimurium is also an open question.

E. coli W and K-12, but not E. coli B and C, mineralize PA (Table 2) through a novel catabolic pathway which does not follow the conventional routes for the aerobic catabolism of aromatic compounds (such as those of E. coli for HPA, 3HPP, and PP degradation) and whose first step is the activation of PA to phenylacetyl-coenzyme A (PA-CoA) by the action of a PA-CoA ligase (94, 328, 182) (Fig. 5B). The second step of the PA catabolic pathway will probably involve the hydroxylation of PA-CoA followed by cleavage of the aromatic ring and further degradation of the resulting aliphatic compound through a -oxidation-like pathway (Fig. 5B). Since the participation of CoA ligases in the initial step of the catabolism of aromatic compounds is a typical feature of anaerobic catabolism (131), the aerobic PA degradation in E. coli constitutes one of the few examples of aerobic hybrid pathways (94, 182).

View larger version (15K): [in this window] [in a new window]   FIG. 5.   Pathway for the catabolism of PA in E. coli. (A) Genetic map of the chromosomal paa cluster. Relevant genes are indicated by blocks: genes with similar shading participate in the same enzymatic step or in the same functional unit of the pathway. Genes flanking the paa cluster (maoA and ydbC) are represented by thick lines. The arrows show the directions of gene transcription. Bent arrows represent the Pz, Pa, and Px promoters. The locations of the IS2 and IS30 insertion sequences within ydbA in E. coli K-12 are shown. The regulatory gene (paaX) is represented by a black block. The inactive and active forms of the PaaX repressor are indicated by empty and solid squares, respectively.  and + indicate transcriptional repression and activation, respectively. The inducer molecule (PA-CoA) is represented by a solid circle. (B) Biochemistry of the PA catabolic pathway. The first intermediate of the pathway is PA-CoA (phenylacetyl CoA). The enzymes are PaaK (PA-CoA ligase), PaaABCDE (putative multicomponent oxygenase), PaaZ (putative ring-cleavage enzyme), and PaaFGHIJ (putative -oxidation-like enzymatic system).

The paa genes responsible for PA degradation in E. coli K-12 and E. coli W strains are organized as a chromosomal cluster of about 14 kb (Fig. 5A), which is absent in the PA-deficient E. coli C strain. However, the ability of E. coli K-12 to grow on PA is strain dependent, with point mutations or small gene rearrangements in the paa cluster being the most probable reason for the PA-deficient phenotype of some K-12 laboratory strains (94) (Table 2).

The E. coli paa cluster contains 14 genes organized in three transcriptional units; two of them, paaZ and paaABCDEFGHIJK, encode the catabolic genes, and the third, paaXY, contains the paaX regulatory gene. All the genes are transcribed in the same direction with the sole exception of paaZ (Fig. 5A). Located downstream of paaZ, paaK, and paaY, three inverted-repeat sequences may act as transcriptional terminators (94). Although the paaK gene product catalyzes the first enzymatic step of the PA catabolic pathway, paaK is located at the 3' end of the paa operon (Fig. 5A), a gene arrangement that explains why Tn1000 insertions within the paa catabolic operon caused polar effects leading to E. coli mutants that did not attack PA (94). The paa genes map at the right end of the mao cluster (Fig. 5A), which is involved in the transformation of PEA into PA (see below), at 31.1 min near the replication terminus of the chromosome (94). This location confirms previous observations that mapped the mutations in two PA-deficient mutants of E. coli K-12 in this chromosomal region (48). Although the left end of the paa cluster is adjacent to the maoA gene both in E. coli W and K-12, the right end of this cluster differs in these two strains. Thus, while the paaY stop codon was found 231 bp upstream of the ATG start codon of the ydbC gene in E. coli W (94), a 9.2-kb sequence containing a long open reading frame (ydbA) disrupted by two insertion sequences (IS2D and IS30C) was found between paaY and ydbC in E. coli K-12 (EcoGene database at the Colibri website) (Fig. 5A). The presence of insertion sequences near the paa cluster and the location of this cluster in a nonessential region of the chromosome (135) provide some clues to the possible mechanisms of gene mobilization of a catabolic cassette and could explain the heterogeneity observed among different E. coli strains to mineralize PA (see below).

Growth of E. coli on PEA as the sole carbon and energy source induces two enzymatic activities that constitute the upper pathway for the catabolism of aromatic amines: an amine oxidase (MaoA) that converts PEA into phenylacetaldehyde (PAL) and a phenylacetaldehyde dehydrogenase (PadA or FeaB) that oxidizes the latter to PA (228) (Fig. 6B). E. coli mutants defective in PadA cannot grow on PEA as a carbon source but can still use it as a nitrogen source (228). Tyramine and dopamine are also substrates of MaoA and PadA, leading to formation of the corresponding aromatic acids, i.e., 4HPA and HPC, respectively (228) (Fig. 6B). Therefore, while E. coli K-12, which lacks the hpa cluster (see above), is able to use tyramine and dopamine only as nitrogen sources, E. coli W, which catabolizes 4HPA and HPC, can use these two aromatic amines as both carbon and nitrogen sources. The reaction catalyzed by MaoA leads to the formation of H₂O₂; therefore, to avoid the toxic effects of the latter, catalase is also induced within E. coli cells growing in the presence of aromatic amines (228).

View larger version (29K): [in this window] [in a new window]   FIG. 6.   Upper pathway for the catabolism of aromatic amines (PEA, tyramine, and dopamine) in E. coli. (A) Genetic map of the chromosomal cluster for the initial catabolism of aromatic amines. Relevant genes are indicated by blocks. Alternative gene names are in parentheses. The paaZ gene from the paa cluster (see Fig. 5) is indicated by a thick line. The arrows show the directions of gene transcription. Bent arrows represent the PmaoB PmaoA and Ppad promoters. The regulatory gene maoB (feaR) is shown by a solid block. The inactive and active forms of the MaoB activator are represented by empty and solid hexagons, respectively. + indicates transcriptional activation. The inducer molecule (PEA, tyramine) is represented by a solid circle. (B) Biochemistry of the initial catabolism of aromatic amines. The metabolites are PAL (phenylacetaldehyde), 4HPAL (4-hydroxyphenylacetaldehyde), DHPAL (3,4-dihydroxyphenylacetaldehyde), PA, 4HPA, and HPC (homoprotocatechuate). The enzymes are MaoA (monoamine oxidase) and PadA (phenylacetaldehyde dehydrogenase).

Expression of the maoA and padA genes is controlled by the MaoB (FeaR) regulator. The maoA, padA, and maoB genes (alternative gene names, tynA, feaB, and feaR, respectively) have been sequenced in E. coli K-12 and W strains and are located at 31.1 min on the chromosome (94, 127, 307). The padA gene is transcribed in the opposite direction to that of maoA and maoB (Fig. 6A), indicating that these three genes, although involved in the same metabolic pathway and physically associated, do not constitute an operon (95). Since these genes map immediately adjacent to the left end of the paa cluster (Figs. 5A and 6A), this supraoperonic clustering of two related pathways, i.e., the upper pathway for PEA catabolism and the PA catabolic pathway, represents an example of physical linkage within the PA-CoA catabolon (182, 217).

The initial step in the aerobic catabolism of 4HPA/3HPA, 3HPP/3HCI, and PP in E. coli is carried out by oxygenases that introduce one (monooxygenases) or two (dioxygenases) atoms of molecular oxygen into the phenyl ring. These oxygenases belong to three different families of ring-hydroxylating oxygenases (Table 3).

4HPA/3HPA monooxygenase. The hpaBC genes located at the 3' end of the hpa cluster in E. coli W encode the two-component monooxygenase that initiates the catabolism of 4HPA and 3HPA in this strain (251) (Fig. 1). This enzyme is a member of the two-component nonheme flavin diffusible monooxygenases (TC-FDM) family, which can be defined according to the following properties: (i) the reductase and the oxygenase components are encoded by two different genes; (ii) the reductase component uses NAD(P)H to catalyze the reduction of a flavin that diffuses to the oxygenase component for oxidation of the substrate by molecular oxygen; and (iii) the two components are not flavoproteins and lack typical ferredoxin and/or flavin/NAD(P)H binding motifs (104).

3HPP monooxygenase. The first catabolic gene of the mhp cluster, that is, mhpA, encodes the MhpA monooxygenase, which hydroxylates 3HPP and 3HCI to DHPP and DHCI, respectively (36, 39) (Figs. 2 and 3). MhpA shows similarity to typical monocomponent bacterial flavin-type aromatic hydroxylases (Table 3). In these proteins, there are three regions of special relevance involved in binding to the FAD molecule: (i) the NH₂ terminus contains the putative fold consensus fingerprint sequence, which is involved in binding of the ADP moiety of FAD; (ii) middle of the protein contains the A(C)DG motif; and (iii) the COOH terminus contains a stretch of amino acids that are likely to participate in binding to the flavin moiety of FAD (75, 92). Although MhpA requires NADH for its activity (36), no NADH binding site could be inferred from the analysis of its amino acid sequence, a characteristic also observed with other flavin monooxygenases (75). Two 3HPP monooxygenases that show similarity to MhpA have been described so far, i.e., HppA from Rhodococcus globerulus PWD1 (18) and MhpA from Comamonas testosteroni TA441 (10) (Table 3). The HppA enzyme appears to have a relatively broad substrate specificity, although PP, 2HPP, 2HCI, and CI were not oxygenated (18). 2HPP (melilotate) is also not oxidized by MhpA, which explains why this aromatic acid cannot be used as carbon source by E. coli (36). Melilotate is the substrate recognized by the flavoprotein melilotate hydroxylase isolated from Pseudomonas spp., an enzyme that does not recognize 3HPP (302). A second melilotate hydroxylase, OhpB, has been described in Rhodococcus sp. strain V49. Although the OhpB enzyme also oxidizes 2HCI, it has low activity on 3HPP, 3HCI, and CI (247). The different substrate specificities of the 3HPP monooxygenases (MhpA and HppA) and the 2HPP monooxygenase (OhpB) reflect their low amino acid sequence similarity (Table 3).

PP dioxygenase. Sequence comparison analyses of the hcaE (formerly hcaA1), hcaF (formerly hcaA2), hcaC, and hcaD gene products revealed significant similarities to the corresponding four protein subunits of the three-component class IIB ring-activating dioxygenases (41) (Table 3). The hcaEFCD genes are suggested, therefore, to encode the HcaEFCD initial dioxygenase of the PP catabolic pathway. This enzyme, together with the HcaB dihydrodiol dehydrogenase, is able to oxidize PP and CI to DHPP and DHCI, respectively (71) (Fig. 4B).

The reaction catalyzed by the ring-opening enzymes metabolizing catecholic compounds is a critical step in the bacterial assimilation of aromatic compounds (129). Thus far, only two ring cleavage dioxygenases have been identified in the catabolism of aromatic compounds in E. coli, i.e., MhpB for DHPP and DHCI cleavage in the PP/3HPP and 3HCI pathways, respectively (Fig. 2 and 3), and HpaD (HpcB) for HPC cleavage in the HPA pathway (Fig. 1). Both enzymes cluster within the type II (87) or class III (298) extradiol dioxygenases, although they belong to two different subfamilies. The HpaD subfamily shows lower similarity among its members, which, in addition, have a deletion of about 40 residues in the central region of their primary structures (175). The two histidines and one glutamate (or aspartate) that form the iron coordination sphere in the class III extradiol dioxygenase whose three-dimensional structure is known, i.e., the LigAB protocatechuate 4,5-dioxygenase from Sphingomonas paucimobilis SYK-6 (306), are conserved in MhpB (His-10, His-53, and Glu-271) and HpaD (His-12, His-57, and Asp-257). Moreover, the histidine residue that functions as the catalytic base (306) is also conserved in MhpB (His-179) and HpaD (His-185).

The hpaD gene from E. coli W encodes a protein of 32 kDa (283 aa) homologous to the HPC 2,3-dioxygenase (276 aa) of E. coli C encoded by the hpcB gene (268). The most relevant difference between the two enzymes was observed in their C-terminal ends, where a 2-bp insertion produces a premature termination in HpcB. Another HPC 2,3-dioxygenase (HpaB) has been described in K. pneumoniae M5a1 (GenBank accession no. AJ000054) and shows 87% amino acid sequence identity to its orthologous enzymes HpaD and HpcB of E. coli. Putative HPC 2,3-dioxygenases that show 93, 85, and 65% identity to those of E. coli are also encoded in the putative hpa clusters of Salmonella species, Y. pestis, and P. aeruginosa, respectively (see below). Both HpcB from E. coli C (268) and the equivalent HpaB protein from K. pneumoniae M5a1 (113) are highly unstable enzymes, and the addition of reagents such as glycerol, acetone, dithiothreitol, or iron salts, which are known to stabilize some ring cleavage dioxygenases, had no effect on these HPC dioxygenases. Unexpectedly, the HPC dioxygenase from K. pneumoniae appears to contain Mg²⁺ instead of Fe²⁺ in its tetrameric structure (113). Whether the HPC 2,3-dioxygenase from E. coli also requires Mg²⁺ for its activity or requires Fe²⁺ (like most extradiol dioxygenases) or Mn²⁺ (like the HPC 2,3-dioxygenase from Arthrobacter globiformis [333]) remains to be determined.

The MhpB dioxygenase responsible for the meta cleavage of DHPP was unstable in cellular crude extracts but could be substantially stabilized by addition of ethanol, glycerol, and iron(II) to the purification buffer (34). The purified protein could be fully reactivated by treatment with iron(II) and a reducing agent such as ascorbate. Therefore, as was found for most of the extradiol dioxygenases, MhpB requires a nonheme Fe²⁺ cofactor for activity and exists as a tetramer (34). The extradiol oxidative cleavage is a complex multistep reaction that highlights the multifaceted catalytic properties of nonheme iron(II): activation of dioxygen, activation of the aromatic ring, carbon-carbon bond cleavage, and insertion of both atoms of molecular oxygen. Semiquinone and lactone intermediates have been identified in the MhpB-mediated extradiol cleavage of DHPP (35). Chemical modification with group-specific reagents revealed that the MhpB enzyme was inactivated by treatment with the histidine-directed reagent diethyl pyrocarbonate, which is in agreement with the identification of conserved histidine residues that may function as iron(II) ligands in the primary structure of MhpB (see above) (298). p-Hydroxymercuribenzoate and dithionitrobenzoic acid, two bulky cysteine-directed reagents, also inactivated MhpB, suggesting that a cysteine residue is present at or near the active site of the enzyme (298).

The MhpB dioxygenase shows a broad specificity toward 3-substituted catechols, with the optimum side chain being that of propionate. Catechol and 4-methylcatechol were also cleaved by MhpB. The methyl esther of the natural substrate, DHPP, was processed at almost identical rates, implying that the carboxylate group is not bound through an electrostatic interaction at the respective active sites of MhpB. Similarly, both 3-ethylcatechol and 3-propylcatechol were processed efficiently by MhpB. The importance of the catecholic hydroxyl groups was confirmed by the observation that neither 2,3-dimethoxyphenylpropionic acid nor 2-aminophenol was accepted as a substrate. Although MhpB processed 3-phenethylcatechol, 2,3-dihydroxybenzoic acid was not used as a substrate (298). DHCI was accepted as a good substrate (Fig. 3), indicating that the enzyme is able to bind the alkyl side chain in a transoid conformation. At high substrate concentrations, substrate inhibition was observed; however, at low substrate concentrations, a sigmoidal kinetics was detected, suggesting cooperativity between protein subunits (298). A similar substrate specificity pattern was observed for the MpcI dioxygenase of Ralstonia eutropha (298), which suggests that this enzyme is also a DHPP dioxygenase. Two other DHPP dioxygenases, MhpB from C. testosteroni TA441 (10) and OhpD from Rhodococcus sp. strain V49 (247), also accept catechol and 3-methylcatechol as substrates. In contrast, the DHPP dioxygenase (HppB) from R. globerulus PWD1 appears to be highly specific for catechols with a relatively large acid group (18). Interestingly, the two DHPP dioxygenases from gram-positive bacteria, i.e., HppB and OhpD, cluster as different subfamilies within the phylogenetic tree of class III extradiol dioxygenases.

As indicated above, following meta cleavage of HPC during the catabolism of 4HPA and 3HPA in E. coli W and C, the resulting CHMS gives rise to succinic semialdehyde and pyruvate through a dehydrogenative route (Fig. 1B).

hpaE and hpcC encode the CHMS dehydrogenases of E. coli W (250) and E. coli C (271), respectively. These two proteins show 98% amino acid sequence identity and catalyze the dehydrogenation of CHMS to CHM (Fig. 1B). The HpcC enzyme has been purified and appears to be a NAD-dependent homodimer (50, 90). Comparisons of HpaE and HpcC with proteins of the aldehyde dehydrogenase superfamily revealed all the motifs that characterize these enzymes (237). Moreover, about 40% identity was observed to other members of the hydroxymuconic semialdehyde dehydrogenase family such as the DmpC and XylG proteins from the phenol pathway of Pseudomonas sp. strain CF600 (249) and the toluene pathway of P. putida (129), respectively (250).

The protein encoded by the hpaF and hpcD genes catalyzes the isomerization of CHM to OPET in E. coli W and C, respectively (267) (Fig. 1B). The HpcD isomerase is a trimeric enzyme with a subunit size of 14 kDa (126 aa). The crystal structure of HpcD has been determined, and a large pocket that is proposed to be the active site was identified. The catalytic mechanism involves a single catalytic base that is the N-terminal proline residue. Two arginine residues interact with the substrate in the active-site region (304, 312). Although the HpcD isomerase catalyzes a reaction analogous to that catalyzed by the 4-oxalocrotonate tautomerase of the meta-cleavage pathway of phenol and toluene on a chemically similar substrate, the two enzymes do not have any apparent sequence similarity. However, comparison of the overall three-dimensional structures of the two enzymes revealed similarities in the catalytic pocket, which confirms that the same catalytic mechanism is operative in both systems. Nevertheless, the two isomerases are specific, and each shows a preference for its own substrate. This substrate specificity in the isomerases seems to be determined by steric constraints in the catalytic cleft (304, 312).

The first catabolic gene of the HPC operon, hpaG in E. coli W (Fig. 1), encodes a protein of 46.9 kDa (429 aa), almost identical to the hpcE gene product from E. coli C (269). However, HpaG is 24 aa longer than HpcE due to a 7-bp deletion at the 3' end of hpcE that produces a premature termination (250). HpcE was reported to be a bifunctional decarboxylase/isomerase monomeric enzyme that catalyzes the magnesium-dependent decarboxylation of OPET to 2-oxo-hept-3-ene-1,7-dioic acid (OHED) through HHDD as an intermediate (Fig. 1B) (108, 145, 269). This type of bifunctional decarboxylase/isomerase activity has not been detected in other aromatic catabolic pathways where the enol-keto isomerization appears to be a dispensable enzymatic step and the actual substrate for the next enzyme in the pathway, a hydratase, is the enol form instead of the keto form (128). Since there are no conclusive data to assume that OHED is the substrate of the HpaH hydratase, it is possible that transformation of HHDD into HHED might occur without a previous enzymatic isomerization step (250) (Fig. 1B). The HpaG and HpcE enzymes may have arisen from a gene duplication event since their N-terminal halves are very similar to their C-terminal ones (269). Reinforcing this assumption, in the genomes of Y. pestis, K. pneumoniae, and P. aeruginosa, the putative HPC meta-cleavage operon contains two hpaG-like genes in tandem (hpaG1 and hpaG2), which through gene fusion might have evolved as the two halves of hpaG in E. coli and Salmonella species (see below). It is worth mentioning that while HpaG (HpcE) shows low identity (below 20%) to other aromatic decarboxylases, such as DmpH and XylI for phenol and toluene catabolism, respectively (250), it shows a significant similarity to fumarylacetoacetate hydrolase that catalyzes the last step in the homogentisate pathway for the catabolism of PA and phenylalanine in Aspergillus nidulans (91) and to fumarylpyruvate hydrolases that catalyze the last step in gentisate degradation (329, 351).

The hpaH gene from E. coli W codes for a protein of 267 aa, whose primary structure shows 94% amino acid sequence identity to that of HpcG (264 aa), the hydratase from E. coli C (97, 271) (Fig. 1). The HpcG enzyme requires metal ions for its activity, and the native protein seems to be an hexamer (97). Although the substrate of HpaH (HpcG) is a dioic acid with two carbon atoms more than the 2-hydroxy-penta-2,4-dienoic acid (HPDA), i.e., the substrate of hydratases of the catechol or catechol-like meta-cleavage pathways such as that of 3HPP (Fig. 2B), there is a significant amino acid sequence identity between HpaH (HpcG) and the hydratases of the latter pathways. For instance, the MhpD hydratase from the 3HPP degradation pathway of E. coli (see below) shows 34% amino acid sequence identity to HpaH, which suggests that they have the same ancestral source. Moreover, as previously reported for the hydratases of catechol meta-cleavage pathways, the HpaH (HpcG) protein presents a relevant similarity to aromatic decarboxylases (250), reinforcing the previous suggestion about the common origin for the hydratases/decarboxylases involved in catabolism of aromatic compounds (269, 271).

The next enzyme of the HPC meta-cleavage dehydrogenative route, the HHED aldolase, is encoded by the last gene in the HPC operon, i.e., hpaI in E. coli W and hpcH in E. coli C (Fig. 1) (250, 303). Unlike the situation mentioned above for the aromatic hydratases, there is no striking identity between HHED aldolase (262 aa) and most of the aldolases of catechol meta-fission pathways. However, HpaI (HpcH) aldolase shows significant amino acid sequence identity (57%) to the BphF aldolase of the catechol meta-cleavage pathway for biphenyl/polychlorinated biphenyl degradation in Rhodococcus sp. strain RHA1. Moreover, since the HpaH (HpcG) hydratase (see above) also reveals a significant similarity to the corresponding BphE hydratase from strain RHA1, the RHA1 meta-cleavage pathway genes may have evolved from the same ancestor as the HPC meta-cleavage pathway genes (189). Two other putative aldolases that show significant similarity to HpaI were found encoded in the genome of Deinococcus radiodurans R1 (331) and in the 184-kb catabolic plasmid of Sphingomonas aromaticivorans F199 (266). Interestingly, two E. coli proteins, YfaU (267 aa) and YhaF (256 aa), show 56 and 48% amino acid sequence identity to HpaI, respectively, suggesting that their genes might encode aldolases that act on substrates chemically similar to HHED. The genes encoding such putative aldolases are located at 50.7 min (yfaU) and 70.5 min (yhaF) on the E. coli K-12 map, and in both cases they are followed by a gene, yfaV and yhaU, respectively, encoding a putative transporter of the anion:cation (H⁺ or Na⁺) symporter (ACS) family of transport proteins (223). A similar gene arrangement is present in the hpa cluster from E. coli W; i.e., the hpaI gene encoding HHED aldolase is followed by hpaX encoding the HPA transporter, also a member of the ACS family of transport proteins (see below) (Fig. 1). An aldol cleavage reaction that resembles that on HHED is found in the catabolism of glucarate and galactarate, and therefore the YhaF protein has been suggested to be the -dehydro--deoxy-D-glucarate aldolase that produces pyruvate and tartronic semialdehyde (151, 303), with the YhaU protein being a putative glucarate transporter (223).

Of the two products generated by the action of HpaI (HpcH) on HHED, i.e., pyruvate and succinic semialdehyde (Fig. 1B), the latter needs to be oxidized to succinic acid before it can enter the Krebs cycle. E. coli strains have two succinic semialdehyde dehydrogenases, a NADP-dependent enzyme encoded by the gabD gene and a NAD-dependent enzyme encoded by a gene (sad) that was mapped at 34 min and has been characterized yet (184, 262, 295). While the GabD enzyme is induced by growth on -aminobutyrate, the NAD-dependent succinic semialdehyde dehydrogenase is induced by growth of the cells in 4HPA or succinic semialdehyde. A NAD-linked succinic semialdehyde dehydrogenase participating in the catabolism of 4HPA in K. pneumoniae has also been described (277). The NAD-dependent enzyme from E. coli is a dimer with a subunit size of 55 kDa (77). E. coli C sad mutants were unable to grow on 4HPA, and this defect was repaired by replacing the affected gene with its orthologous gene from E. coli K-12 (295). Since the sad gene is not within the hpa cluster and is present in E. coli K-12 strains, which lack the 4HPA pathway, it is likely that the NAD-dependent succinic semialdehyde dehydrogenase may fulfill different physiological roles in E. coli, such as preventing the toxic effect of the succinic semialdehyde generated when the cells grow on -aminobutyrate as the sole carbon and nitrogen source (77).

During the catabolism of 3HPP and PP in E. coli, ring cleavage of DHPP by the MhpB dioxygenase (see above) give rise to 2-hydroxy-6-keto-nona-2,4-diene 1,9 dioic acid (HKNDA) (Fig. 2B). Further catabolism of HKNDA through a hydrolytic route ends with the formation of succinic acid, pyruvic acid, and acetyl-CoA, which enter the Krebs cycle (36, 39) (Fig. 2).

The mhpC gene encodes the HKNDA hydrolase (MhpC), which catalyzes the hydrolytic cleavage of the extradiol ring fission product of DHPP, giving rise to succinic acid (which enters the Krebs cycle) and HPDA (Fig. 2B). The purified MhpC protein (287 aa) lacks the initial methionine residue and is a homodimer that does not require cofactors for its catalytic activity (170). In contrast to the ring cleavage dioxygenase MhpB (see above), the MhpC hydrolase shows some substrate selectivity for the carboxylate of the side chain. Slow turnover was observed for the ring fission products of 3-methylcatechol or catechol. However, the ring fission product of DHCI was a fairly efficient substrate, generating fumaric acid and HPDA (170) (Fig. 3). A single active site is responsible for the enzymatic reaction, which involves a fast initial ketonization of the dienol substrate followed by a stereospecific C-C fragmentation step. The dissociation of the keto intermediate from the enzyme prior to hydrolysis explains the uncoupling of substrate utilization and product formation (134, 169, 170). This leakiness has been also found in other C-C bond hydrolases for the catabolism of