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Metabolic reconstruction of aromatic compounds degradation from the genome of the amazing pollutant-degrading bacterium Cupriavidus necator JMP134

Danilo Pérez-Pantoja, Rodrigo De la Iglesia, Dietmar H. Pieper, Bernardo González
DOI: http://dx.doi.org/10.1111/j.1574-6976.2008.00122.x 736-794 First published online: 1 August 2008


Cupriavidus necator JMP134 is a model for chloroaromatics biodegradation, capable of mineralizing 2,4-D, halobenzoates, chlorophenols and nitrophenols, among other aromatic compounds. We performed the metabolic reconstruction of aromatics degradation, linking the catabolic abilities predicted in silico from the complete genome sequence with the range of compounds that support growth of this bacterium. Of the 140 aromatic compounds tested, 60 serve as a sole carbon and energy source for this strain, strongly correlating with those catabolic abilities predicted from genomic data. Almost all the main ring-cleavage pathways for aromatic compounds are found in C. necator: the β-ketoadipate pathway, with its catechol, chlorocatechol, methylcatechol and protocatechuate ortho ring-cleavage branches; the (methyl)catechol meta ring-cleavage pathway; the gentisate pathway; the homogentisate pathway; the 2,3-dihydroxyphenylpropionate pathway; the (chloro)hydroxyquinol pathway; the (amino)hydroquinone pathway; the phenylacetyl-CoA pathway; the 2-aminobenzoyl-CoA pathway; the benzoyl-CoA pathway and the 3-hydroxyanthranilate pathway. A broad spectrum of peripheral reactions channel substituted aromatics into these ring cleavage pathways. Gene redundancy seems to play a significant role in the catabolic potential of this bacterium. The literature on the biochemistry and genetics of aromatic compounds degradation is reviewed based on the genomic data. The findings on aromatic compounds biodegradation in C. necator reviewed here can easily be extrapolated to other environmentally relevant bacteria, whose genomes also possess a significant proportion of catabolic genes.

  • metabolic reconstruction
  • aromatic compounds
  • degradation
  • Cupriavidus necator


Cupriavidus necator JMP134 (ex Alcaligenes eutrophus; ex Ralstonia eutropha; ex Wautersia eutropha) was isolated from an Australian soil by its ability to grow on 2,4-dichlorophenoxyacetate (2,4-D) (Pemberton, 1979; Don & Pemberton, 1981). Early studies also showed that this strain grows on 4-methyl-2-chlorophenoxyacetate (MCPA) and 3-chlorobenzoate (3-CB) (Pemberton, 1979; Don & Pemberton, 1981), and is resistant to mercurial compounds. The determinants for 2,4-D and 3-CB degradation and for mercury resistance are encoded in the catabolic plasmid pJP4 (Don & Pemberton, 1981). The presence of the tfd genes on this plasmid, which encode the ortho ring-cleavage pathway for chlorocatechol intermediates, facilitated the use of this bacterium as a model for chloroaromatic degradation. A number of articles have dealt with the presence of tfd and tfd-like sequences both in pristine and in polluted environments, and tfd genes have also been used to track the presence and transference of the pJP4 plasmid and related plasmids in the environment. These topics have been covered in a couple of reviews (Top, 1998, 2002).

In recent years, the discovery of a second pJP4-encoded tfd genes cluster has led to a revision of the role of the tfd-encoded functions (Leveau, 1999; Laemmli, 2000, 2004; Perez-Pantoja, 2000; Plumeier, 2002; Schlomann, 2002). In addition, increasing evidence indicates that C. necator JMP134 grows on several other pollutants and natural compounds using chromosomally encoded functions (Schlomann, 1990b; Ecker, 1992; Schenzle, 1997; Padilla, 2000; Louie, 2002). The recent availability of the complete genome sequence of several environmentally relevant bacteria such as Pseudomonas putida KT2440 (Nelson, 2002); Burkholderia xenovorans LB400 (Chain, 2006); Rhodococcus sp. RHA1 (McLeod, 2006), and the denitrifying bacterium ‘Aromatoleum aromaticum’ sp. strain EbN1 (Rabus, 2005), has revealed the presence of a significant number of genes encoding determinants for aromatic compounds degradation. This fact suggests that different bacteria are potentially able to degrade several types of both natural and man-made aromatic compounds. The genome sequence of C. necator also reveals the presence of a significant (more than 5% of all the putative genes) proportion of aromatics degradation genes. We selected this versatile pollutant-degrading bacterium as a model of aromatic compounds degraders, and performed the metabolic reconstruction of aromatic compounds degradation. In this review, we analyze the main features of the catabolism of aromatic compounds in C. necator JMP134 within the context of the abundant literature on this topic. Special attention is given to those aspects that would explain the impressive catabolic versatility of this and other bacteria.

Aromatic growth substrates for C. necator JMP134

Out of 140 aromatic substrates tested, 60 can be used by C. necator JMP134 as the sole carbon and energy source; these include c. 40 compounds that have not been reported previously as growth substrates. The growth supporting aromatic compounds are (brackets indicate the section where the corresponding degradation pathway is discussed): benzoate, benzaldehyde, benzyl alcohol, phenylglyoxylate (benzoylformate), benzyl acetate, benzylamine (‘The cat and ben genes’, benzoate also in ‘The aerobic benzoyl-CoA pathway’); 4-hydroxybenzoate, 3,4-dihydroxybenzoate (protocatechuate), chlorogenate, quinate (‘The pob and pca genes’); phenylpropionate, cinnamate, 4-hydroxyphenylpropionate, 4-hydroxycinnamate (coumarate), 3,4-dihydroxyphenylpropionate, 3,4-dihydroxycinnamate (caffeate), ferulate (‘Peripheral pathways that channel phenylpropenoid and phenylpropanoid compounds to the β-ketoadipate pathway’, and some of these compounds in ‘The 2,3-dihydroxyphenylpropionate meta ring-cleavage pathway’); 3-hydroxyphenylpropionate (‘The 2,3-dihydroxyphenylpropionate meta ring-cleavage pathway’); 2-hydroxy (salicylate), 3-hydroxybenzoate, 3-hydroxybenzyl alcohol, 2,5-dihydroxybenzoate (gentisate), ethylsalicylate (‘Degradation of salicylate and 3-hydroxybenzoate: the gentisate pathway’); phenol, 2-, 3-, and 4-methylphenol, 2,3- and 3,4-dimethylphenol, 4-ethylphenol, 2-methylphenoxyacetate, benzene, toluene [‘The catabolic pathways for benzene, toluene and (methyl)phenols’]; phenylacetate, phenylacetaldehyde, phenylethylamine, phenylpyruvate, 4-phenylbutyrate, 5-phenylvalerate, 6-phenylhexanoate (‘The phenylacetyl-CoA ring-cleavage pathway’); 2-, 3-, and 4-hydroxyphenylacetate, 4-hydroxyphenylpyruvate, phenylalanine, tyrosine (‘The homogentisate ring-cleavage pathway’); tryptophan, 2-aminobenzoate (anthranilate) [‘The 2-aminobenzoyl-CoA pathway’]; 3-hydroxyanthranilate (‘The 3-hydroxyanthranilate pathway’); 3-nitrophenol, 2-chloro-5-nitrophenol (‘Catabolic pathways for nitrophenols’); hydroquinone, 2,4,6-trichlorophenol [‘The (chloro)hydroxyquinol ring-cleavage pathway’]; 4-fluorobenzoate (‘Fluorobenzoate catabolism’); 3-CB, 2,4-D and MCPA (‘Catabolic pathway for mono- and dichlorinated compounds: the tfd genes’). In addition, strain JMP134 grows on 2,6-dinitrophenol and 3,5-dihydroxybenzoate, although for these compounds no degradation pathway could be identified. Cell yields range from 0.9 to 5.7 mg cells mmol−1 of carbon. Chlorinated compounds, aromatic aldehydes and ferulate produce lower growth yields. Eighty compounds failed to support the growth of this strain: 2- and 4-chlorobenzoate, 3,5-dichlorobenzoate, 3-chloro-4-hydroxybenzoate, 2- and 4-chlorophenoxyacetate, 3-(2,4-dichlorophenoxy)propionate, 4-(2,4-dichlorophenoxy)butyrate, 2-, 3- and 4-chlorophenylacetate, 3-chloro-4-hydroxyphenylacetate, tropate, 2- and 3-phenoxypropionate, 4-phenoxybutyrate, 2-hydroxyphenylpropionate, 2-hydroxycinnamate, 2-phenylpropionate, 2-, 3- and 4-methoxybenzoate, 2,6-, 2,3- and 2,4-dihydroxybenzoate, 3- and 4-methylbenzoate, 3- and 4-phenoxybenzoate, 2- and 4-hydroxybenzylalcohol, 3,5-dihydroxybenzylalcohol, phthalate, 1,2-, 1,3- and 1,4-dimethylbenzene, ethylbenzene, chlorobenzene, biphenyl, coniferyl alcohol, vanillyl alcohol, 4-hydroxy-3-methoxybenzoate (vanillate), 3-hydroxy-4-methoxybenzoate (isovanillate), isovanillyl alcohol, vanillaldehyde, 3-hydroxy-4-methoxycinnamate, 5-chlorovanillate, 2-, 3- and 4-chlorophenol, 2,3-, 2,5-, 2,6-, 3,4- and 3,5-dichlorophenol, 2,3,4-, 2,3,5- and 2,4,5-trichlorophenol, pentachlorophenol, 2,4- and 3,5-dimethylphenol, 4-propylphenol, resorcinol, mandelate, 4-hydroxymandelate, 3-, 4- and 5-chlorosalicylate, 3,5-dichlorosalicylate, 5-methoxysalicylate, syringate, 4-hydroxyacetophenone, acetophenone, 2-, 3- and 4-nitrobenzoate, 2- and 4-nitrophenol, 4-sulfophenol, 4-sulfobenzoate.

The ortho ring-cleavage pathways for benzoate and p-hydroxybenzoate (the β-ketoadipate central pathway)

The β-ketoadipate pathway is widely distributed among soil bacteria because it plays a central role in the degradation of naturally occurring aromatic compounds (Harwood & Parales, 1996). Some environmental pollutants are also degraded through this pathway. The central reactions of the different branches of the β-ketoadipate pathway in C. necator JMP134 are shown in Fig. 1. The catechol branch, encoded by cat genes, converts the catechol generated from benzoate (through the action of the ben gene products), phenol and some lignin monomers, into β-ketoadipate. The protocatechuate branch, encoded by pca genes, converts the protocatechuate derived from 4-hydroxybenzoate (through the action of the pob gene products) and numerous lignin monomers, into β-ketoadipate. Two additional steps accomplish the conversion of β-ketoadipate into the Krebs cycle intermediates: succinyl-CoA and acetyl-CoA (Harwood & Parales, 1996). Biochemical studies and amino acid sequence data indicate that the pathway enzymes are highly conserved among the phylogenetically diverse organisms that possess this pathway. Despite this biochemical conservation, studies of a limited number of soil bacteria demonstrate a remarkable diversity of this pathway in terms of gene organization, type of inducers and regulation mechanism (Harwood & Parales, 1996).

Figure 1

The β-ketoadipate pathway and peripheral reactions. In this and all the equivalent figures, compounds depicted at the beginning of each biochemical pathway are growth substrates for Cupriavidus necator JMP134, and the gene product names for those cases where genetic or biochemical evidence for function is available, are underlined.

The cat and ben genes

The ben and cat gene products of C. necator JMP134 are highly similar, in their amino acid sequence, to proteins of the catechol branch of the β-ketoadipate pathway that has been characterized in other bacteria, mainly Acinetobacter, Pseudomonas, and Burkholderia (Table 1). The gene encoding CatA1, which starts the catechol branch of the β-ketoadipate pathway (Fig. 1), clusters together with the benABCD genes (small chromosome [C2] in Fig. 2), which are responsible for funneling benzoate into this pathway. This putative operon includes the catR1 gene; this gene encodes a putative LysR-type regulatory protein. A gene putatively encoding a muconate cycloisomerase, catB1, is localized close to the ben and catA1 genes, but is not part of this putative operon (Fig. 2, C2). A similar organization of the β-ketoadipate pathway genes has been observed previously in Burkholderia sp. strain TH2 (Suzuki, 2002), Burkholderia sp. NK8 (Francisco, 2001), and C. necator 335 (accession number AF042281, I.S. Hinner et al., unpublished data). However, unlike those strains where a gene encoding muconolactone isomerase is positioned directly downstream of the muconate cycloisomerase encoding gene, no additional β-ketoadipate pathway genes are localized downstream of catB1 in strain JMP134 (Fig. 2, C2). The catC gene is clearly located in a second operon, which also encodes a muconate cycloisomerase and an enol lactone hydrolase, namely enzymes for the conversion of muconate to β-ketoadipate (large chromosome [C1] in Fig. 2). This gene organization contrasts the one previously observed in Pseudomonas strains, where operons encoding cat genes do not comprise a catD gene; it also supports reports suggesting that catechol and protocatechuate branches of the β-ketoadipate pathway in Cupriavidus strains converge at the stage of β-ketoadipate rather than β-ketoadipate enol lactone, as is the case in Pseudomonas strains (Harwood & Parales, 1996; Jimenez, 2002). Genes encoding β-ketoadipyl-CoA transferase and thiolase are not comprised in the ben/cat genes operons (see next section), as opposed to Acinetobacter, where a full set of genes for the transformation of muconate into succinyl-CoA and acetyl-CoA is present in the cat operons (Harwood & Parales, 1996).

View this table:
Table 1

Genes encoding the β-ketoadipate pathway, the methylmuconolactone pathway and peripheral reactions

Related gene products
GenePosition (bp)No. aaNameFunction/descriptionOrganism% Id (aa)Accession no.References
catB1C2 1058420-1057302372catB4Muconate cycloisomeraseCupriavidus necator H1693 (372)CAJ95333Pohlmann (2006)
catR1C2 1059438-1058521309catRPutative regulator of catechol degradationCupriavidus necator 335T84 (307)AAG42034I.S. Hinner et al. (unpublished data)
catA1 C2 1059617-1060540307catACatechol-1,2-dioxygenaseCupriavidus necator 335T82 (309)AAG42033I.S. Hinner et al. (unpublished data)
benA C2 1060637-1062034465benABenzoate-1,2-dioxygenase α subunitAcinetobacter baylyi ADP175 (437)P07769Neidle (1991)
benB C2 1062031-1062522163benBBenzoate-1,2-dioxygenase β subunitAcinetobacter baylyi ADP169 (163)P07770Neidle (1991)
benCC2 1062586-1063605339benCBenzoate-1,2-dioxygenase electron transfer componentAcinetobacter baylyi ADP166 (335)P07771Neidle (1991)
benDC2 1063611-1064396261benDBenzoate cis-diol dehydrogenasePseudomonas putida KT244070 (258)AAN68772Jimenez (2002)
catR2C1 1823192-1822311293catRPositive regulator of cat operonPseudomonas resinovorans CA1061 (289)BAB32455Nojiri (2002)
catB2C1 1823294-1824418374catB4Muconate cycloisomeraseCupriavidus necator H1694 (371)CAJ95333Pohlmann (2006)
catC C1 1824450-182472892catC3Muconolactone δ-isomeraseCupriavidus necator H1686 (92)CAJ93068Pohlmann (2006)
catXC1 1824784-1825779331h16_B0537Probable extra-cytoplasmic solute receptorCupriavidus necator H1692 (331)CAJ95334Pohlmann (2006)
catDC1 1825813-1826622269catD13-Oxoadipate enol-lactone hydrolaseCupriavidus necator 335T69 (257)AAG42037I.S. Hinner et al. (unpublished data)
pobBC2 645961-647148395pobB4-Hydroxybenzoate-3-monooxygenasePseudomonas sp. CBS359 (386)CAA52824Seibold (1996)
pobAC2 1753176-1752007389pobA4-Hydroxybenzoate-3-monooxygenasePseudomonas fluorescens68 (390)CAA48483van Berkel (1992)
pobRC2 1753301-1754188295pobRRegulatory protein, putative pobA regulatorAzotobacter chroococcum ATCC904337 (287)AAF03756Quinn (2001)
pcaLC2 1755385-1754195396pcaC4-Carboxymuconolactone decarboxylaseRalstonia solanacearum GMI100074 (124)CAD15956Salanoubat (2002)
catD13-Oxoadipate enol-lactone hydrolaseCupriavidus necator 335T52 (250)AAG42037I.S. Hinner et al. (unpublished data)
pcaLEnol-lactone hydrolase/4-CML decarboxylaseRhodococcus opacus 1CP42 (380)AAC38246Eulberg (1998)
pcaBC2 1756810-1755422462pcaB13-Carboxy-cis,cis-muconate cycloisomeraseCupriavidus necator H1682 (450)CAJ97071Pohlmann (2006)
pcaGC2 1757440-1756871189pcaGProtocatechuate-3,4-dioxygenase α chainBurkholderia cepacia DBO147 (201)AAA25925Zylstra (1989)
pcaHC2 1758146-1757445233pcaHProtocatechuate-3,4-dioxygenase β subunitBurkholderia cepacia DBO163 (224)AAA25924Zylstra (1989)
pcaQC2 1758242-1759219325pcaQTranscriptional regulator, LysR familyCupriavidus necator H1685 (313)CAJ97074Pohlmann (2006)
pcaKC2 1760586-1759213457pcaK4-Hydroxybenzoate transporterPseudomonas putida PRS200055 (437)AAA85137Harwood (1994)
pcaI C2 2403520-2404179219pcaIβ-Ketoadipate succinyl-CoA transferasePseudomonas putida PRS200074 (219)AAA25922Parales & Harwood (1992)
pcaJ C2 2404196-2404834212pcaJβ-Ketoadipate succinyl-CoA transferasePseudomonas putida PRS200072 (210)AAA25923Parales & Harwood (1992)
pcaF C2 2404863-2406065400pcaFβ-Ketoadipyl-CoA-thiolaseCupriavidus necator H1695 (400)CAJ95001Pohlmann (2006)
hcaCC2 1584218-1582320632fcsFeruloyl-CoA-synthetasePseudomonas sp. HR19963 (579)CAB60226Overhage (1999)
hcaBC2 1585744-1584293483vdhVanillin dehydrogenasePseudomonas sp. HR19967 (483)CAA72286Priefert (1997)
hcaAC2 1586669-1585836277ORFA4-Hydroxycinnamoyl CoA hydratase/lyasePseudomonas fluorescens AN10384 (276)CAA73502Gasson (1998)
hcaXC2 1587919-1586804371RSc1084Probable porin transmembrane proteinRalstonia solanacearum GMI100071 (359)CAD14786Salanoubat (2002)
hcaKC2 1589180-1587966404hcaKPutative hydroxycinnamate transporterAcinetobacter baylyi ADP151 (393)AAP78948Smith (2003)
hcaGC2 1590969-1589206587hcaGChlorogenate esteraseAcinetobacter baylyi ADP153 (583)AAL54855Smith (2003)
hcaRC2 1591203-1591688161hcaRRepressor protein, MarR familyAcinetobacter baylyi ADP146 (133)AAP78949Smith (2003)
mmlJ C1 1622626-162235191mmlJPutative methylmuconolactone isomeraseCupriavidus necator H16 pHG187 (91)AAP86133Schwartz (2003)
mmlI C1 1623007-1622666113mmlIPutative 4-methylmuconolactone methylisomeraseCupriavidus necator H16 pHG183 (112)AAP86134Schwartz (2003)
mmlHC1 1624359-1623073428mmlHPutative muconolactone transporterCupriavidus necator H16 pHG186 (428)AAP86135Schwartz (2003)
mmlGC1 1625108-1624446220catJOxoadipate-CoA transferase β subunitCupriavidus necator H16 pHG189 (220)AAP86136Schwartz (2003)
mmlFC1 1625811-1625113232catIOxoadipate-CoA transferase α subunitCupriavidus necator H16 pHG188 (231)AAP86137Schwartz (2003)
mmlRC1 1626849-1625935304PHG389Putative regulatory protein LysR-typeCupriavidus necator H16 pHG191 (300)AAP86138Schwartz (2003)
mmlLC1 1627858-1626968296PHG390Hypothetical protein (Zn-dependent hydrolases)Cupriavidus necator H16 pHG191 (296)AAP86139Schwartz (2003)
  • * Genes for those cases where genetic or biochemical evidence for function is available are underlined

Figure 2

Position of aromatic degradation genes in the two chromosomes and the megaplasmid of Cupriavidus necator JMP134. C1, large chromosome; C2, small chromosome; pJPL, megaplasmid. Numbers below the scale bars correspond to million of base pairs.

Both catA1benABCD and catB2CXD gene clusters are preceded by putative catR genes that encode LysR-type regulatory proteins. An interesting point for further investigation would be whether both catR genes are required for full induction of the benzoate degradation pathway. Benzoate and muconate have been reported as inducers of the benzoate pathway in other bacteria, through the activity of LysR-type regulatory proteins (Harwood & Parales, 1996; McFall, 1998; Bundy, 2002). It could be speculated that CatR1 gene product is responsive to benzoate and muconate, whereas the CatR2 gene product responds predominantly to the muconate generated by the CatA1 gene product, which indicates a sequential and modular induction of the cat genes, similar to that observed in Acinetobacter baylyi ADP1 (Ezezika, 2006). Further evidence for a modular gene organization is the fact that a second catechol-1,2-dioxygenase encoding gene, catA2 (Fig. 2, C1, and pathway in Fig. 3), is localized in a cluster of genes encoding a multicomponent phenol hydroxylase [see ‘The catabolic pathways for benzene, toluene and (methyl)phenols’]; this enzyme would probably channel at least some phenol when added as growth substrate, into the β-ketoadipate pathway (Pieper, 1989). CatA1 and CatA2 gene products from C. necator cluster together in the dendrogram of the intradiol 1,2-dioxygenases (Fig. 4).

Figure 3

Catabolic pathways for benzene, toluene, (methyl)phenols, (methyl)phenoxyacetate, 3-hydroxyphenylpropionate, 3-hydroxycinnamate and 3-hydroxyanthranilate. R: methyl or H.

Figure 4

Dendrogram showing the relatedness of intradiol-1,2-dioxygenases. The dendrogram was obtained by the neighbor-joining method using mega 4.0 based on sequence alignments calculated by clustal w using the default options. Sequences of deduced proteins encoded in the genome of Cupriavidus necator JMP134 are highlighted black. The sequences and their accession numbers are as follows: TfdCI P4a, Delftia acidovorans P4a chlorocatechol-1,2-dioxygenase (AAC35836); TfdCII P4a, D. acidovorans P4a chlorocatechol-1,2-dioxygenase (AAM76776); TetC RW71, Pseudomonas chlororaphis RW71 chlorocatechol-1,2-dioxygenase II (CAB52140); TcbC P51, Pseudomonas sp. strain P51 catechol-1,2-dioxygenase II (AAD13625); ClcA B13, Pseudomonas knackmussii B13 chlorocatechol-1,2-dioxygenase (CAE92861); TfdCII JMP134, C. necator JMP134 chlorocatechol-1,2-dioxygenase (YP_025394); TfdC 2a, Burkholderia cepacia 2a chlorocatechol-1,2-dioxygenase (AAK81678); TfdC EST4002, Achromobacter xylosooxidans ssp. denitrificans EST4002 chlorocatechol-1,2-dioxygenase (AAS49440); TfdCI JMP134, C. necator JMP134 chlorocatechol-1,2-dioxygenase (YP_025387); TfdC NK8, Burkholderia sp. strain NK8 chlorocatechol-1,2-dioxygenase (BAB56009); TfdC2 tfd44, Sphingomonas sp. strain tfd44 chlorocatechol-1,2-dioxygenase (AAT99373); DccAII MH, Sphingobium herbicidovorans MH chlorocatechol-1,2-dioxygenase (CAF32822); DccAI MH, S. herbicidovorans MH chlorocatechol-1,2-dioxygenase (CAF32818); TfdC tfd44, Sphingomonas sp. strain tfd44 chlorocatechol-1,2-dioxygenase (AAT99364); ClpC S1, Defluvibacter lusatiensis S1 chlorocatechol-1,2-dioxygenase (CAD60254); ClcA1 1CP, Rhodococcus opacus 1CP chlorocatechol-1,2-dioxygenase (AAC38251); ClcA2 1CP, R. opacus 1CP chlorocatechol-1,2-dioxygenase (CAD28142); CatA-II BA-5-17, Arthrobacter sp. strain BA-5-17 catechol-1,2-dioxygenase (BAD11154); CatA mA3, Arthrobacter sp. strain mA3 catechol-1,2-dioxygenase (CAA03944); CatA ATCC 39116, Streptomyces setonii ATCC 39116 catechol-1,2-dioxygenase (AAK14065); CatA 1CP, R. opacus 1CP catechol-1,2-dioxygenase (CAA67941); CatA NCIMB 13259, R. rhodochrous NCIMB 13259 catechol-1,2-dioxygenase (AAC33003); CatA AN-13, Rhodococcus erythropolis AN-13 catechol-1,2-dioxygenase (BAA11859); CatA KT2440, Pseudomonas putida KT2440 catechol-1,2-dioxygenase (NP_745846); CatA2 KT2440, P. putida KT2440 catechol-1,2-dioxygenase (NP_745310); CatA CA10, Pseudomonas resinovorans CA10 catechol-1,2-dioxygenase (BAB32458); SalD MT1, Pseudomonas reinekei MT1 catechol-1,2-dioxygenase (ABH07022); CatA MT1, P. reinekei MT1 catechol-1,2-dioxygenase (ABI93947); PheB EST1001, Pseudomonas sp. strain EST1001 catechol-1,2-dioxygenase (AAC64900); CatA ADP1, Acinetobacter bayly ADP1 catechol-1,2-dioxygenase (YP_046127); IsoA S13, A. radioresistens S13 catechol-1,2-dioxygenase (AAK55425); CatA1 ANA-18, Frateuria sp. strain ANA-18 catechol-1,2-dioxygenase (BAC82534); CatA1 TH2, Burkholderia sp. strain TH2 catechol-1,2-dioxygenase (BAC16779); CatA2 ANA-18, Frateuria sp. strain ANA-18 catechol-1,2-dioxygenase (BAA75211); CatA2 TH2, Burkholderia sp. strain TH2 catechol-1,2-dioxygenase (BAC16769); CatA2 JMP134, C. necator JMP134 catechol-1,2-dioxygenase (YP_295914); CatA NCIB8250, Acinetobacter calcoaceticus NCIB8250 catechol-1,2-dioxygenase (CAA85386); IsoB S13, Actinobacter radioresistens S13 catechol-1,2-dioxygenase (AAG16896); CatA 335, Ralstonia eutropha 335 catechol-1,2-dioxygenase (AAG42033); CatA1 JMP134, C. necator JMP134 catechol-1,2-dioxygenase (YP_298598); ReutB5855 JMP134, C. necator JMP134 intradiol dioxygenase (YP_300042); PnpC SJ98, Ralstonia sp. strain SJ98 hydroxyquinol-1,2-dioxygenase (AAS87586); GraB MTP-10005, Rhizobium sp. MTP-10005 hydroxyquinol-1,2-dioxygenase (BAF44523); DxnF RW1, Sphingomonas sp. strain RW1 catechol-1,2-dioxygenase (ZP_01610139); NCgl1113 ATCC13032, Corynebacterium glutamicum ATCC 13032 hydroxyquinol-1,2-dioxygenase (NP_600386); NCgl2951 ATCC13032, C. glutamicum ATCC 13032 hydroxyquinol-1,2-dioxygenase (NP_602248); ORF2 BA-5-17, Arthrobacter sp. strain BA-5-17 hydroxyquinol-1,2-dioxygenase (BAA82713); NpcC SAO101, R. opacus SAO101 hydroxyquinol-1,2-dioxygenase (BAD30043); TftH AC1100, B. cepacia AC1100 hydroxyquinol-1,2-dioxygenase (AAC43338); HxqC JMP134, C. necator JMP134 hydroxyquinol-1,2-dioxygenase (AAZ60953); ORF3 R34, B. cepacia R34 hydroxyquinol-1,2-dioxygenase (AAL50018); TcpC JMP134, C. necator JMP134 (chloro)hydroxyquinol 1,2-dioxygenase (AAZ63491); HadC DTP0602, Ralstonia pickettii DTP0602 hydroxyquinol 1,2-dioxygenase (BAA13107).

The pob and pca genes

The pca genes that encode enzymes of the protocatechuate branch of the β-ketoadipate pathway in C. necator JMP134 are organized in two clusters: pobAR-pcaLBGHQK and pcaIJF (C2 in Fig. 2), unlike gene organizations described previously. The pcaIJF genes encode the enzymes required for the conversion of β-ketoadipate to the Krebs cycle's intermediates; these catabolic steps are common to both branches of the β-ketoadipate pathway (Fig. 1, Table 1). In several Proteobacteria, β-ketoadipate is the inducer of pcaIJF genes by activation of PcaR/PcaQ, transcriptional regulators of the IclR-type family (Harwood & Parales, 1996). However, pcaR/pcaQ genes are not found in the vicinity of the pcaIJF genes in C. necator JMP134. A gene encoding a putative LysR-type regulator homologous to the pcaR gene of Pseudomonas strains (about a 40% aa identity) is located 9 kb away from the pcaK gene in C. necator JMP134, and could be the regulatory gene involved in pcaIJF gene induction. It is not uncommon that transcriptional regulators control the expression of distal genes. For example, PcaR gene products in P. putida are encoded at a distance from the target pcaHG genes (Harwood & Parales, 1996). Alternatively, the fact that in strain JMP134 the pcaIJF genes are not linked to the pca gene cluster would indicate that these gene functions are involved in other CoA transferase activities, such as those reported in the degradation of straight-chain dicarboxylic acids (Parke, 2001); therefore, they would be controlled by different regulatory proteins and/or additional inducers when nonaromatic substrates are metabolized through the activities of the PcaIJF gene products. The pca gene organization in C. necator JMP134 is different from that found in Ralstonia solanacearum (two clusters), P. putida (four clusters) and A. baylyi. ADP1 (one supraoperonic cluster). There exists, then, a great diversity in the genetic organization of this pathway in Proteobacteria (Harwood & Parales, 1996). Furthermore, gene order does not appear to be maintained within the clusters, except in cases where genes may coevolve because they encode subunits of a single enzyme and are cotranscribed; e.g., pcaGH genes encoding protocatechuate dioxygenase and pcaIJ genes encoding β-ketoadipate succinyl-CoA transferase (Harwood & Parales, 1996). A striking aspect of the pca genes in C. necator JMP134 is that pcaC and pcaD genes – encoding γ-carboxymuconolactone decarboxylase and β-ketoadipate enol-lactone hydrolase, respectively, two enzymes that perform successive steps in the protocatechuate degradation branch (Fig. 1) – are fused in a unique pcaL gene (C2 in Fig. 2). Sequence analysis of pcaL gene reveals that the N-terminal two thirds of the protein are homologous to the enol-lactone hydrolases, whereas the C-terminal third is homologous to the decarboxylases (Table 1). Such a gene fusion is not observed in R. solanacearum, P. putida and Bradyrhizobium japonicum, where pcaD and pcaC genes are located together (Lorite, 1998; Jimenez, 2002; Salanoubat, 2002), in what seems to be a previous stage of the gene fusion in C. necator JMP134. A similar gene fusion of pcaD and pcaC genes has been described previously in Rhodococcus opacus 1CP (Eulberg, 1998), in Streptomyces sp. strain 2065 (Iwagami, 2000) and, very recently, in Acinetobacter baumannii DU202 (Park, 2006). Gene databases also show the presence of similar gene fusions in unrelated bacteria such as Caulobacter, Nocardioides and Mycobacterium (data not shown). Sequence comparison of pcaL genes indicate that the gene fusions in C. necator and R. opacus took place separately, and are not due to a horizontal gene transfer from Gram-positive bacteria to C. necator, because each catalytic domain in the fused PcaL gene product of C. necator has a much higher identity with the proteobacterial PcaC and PcaD counterparts than with the PcaL gene product from R. opacus (Table 1). The fact that these gene fusions are present in distantly related bacterial groups strongly suggests a biochemical advantage of these fused gene products.

Another striking aspect of the protocatechuate branch genes in C. necator JMP134 is the presence of two genes, pobA and pobB that putatively encode p-hydroxybenzoate hydroxylases. The position of both gene products in the dendrogram of FAD-dependent hydroxylases is shown in Fig. 5. The pobA and pobB genes in C. necator JMP134 have a 59% aa identity, indicating a rather far evolutionary origin. However, both gene products contain the sequence Gly–X–Gly–X–X–Gly (residues 9–14), which is characteristic of flavoproteins. These glycine residues have been claimed to play an important structural role (Hofsteenge, 1980). The first case in which two genes encode a p-hydroxybenzoate hydroxylase has been reported in Pseudomonas fluorescens, in which an isoenzyme gene was cloned and showed to express half of the total p-hydroxybenzoate hydroxylase activity (Shuman & Dix, 1993). However, this isoenzyme was not expressed during growth of P. fluorescens on 4-hydroxybenzoate. It could be of interest to investigate whether pobA and pobB genes are differential or simultaneously expressed in C. necator JMP134.

Figure 5

Dendrogram showing the relatedness of FAD-dependent monooxygenases. The dendrogram was obtained by the neighbor-joining method using mega 4.0 based on sequence alignments calculated by clustal w using the default options. Sequences of deduced proteins encoded in the genome of Cupriavidus necator JMP134 are highlighted black. The sequences and their accession numbers are as follows: TfdB P4a, Delftia acidovorans P4a dichlorophenol hydroxylase (AAM76774); TfdB EST4002, Achromobacter xylosoxidans ssp. denitrificans; TfdBII JMP134, C. necator JMP134 2,4-dichlorophenol hydroxylase (YP_025391); TfdBI JMP134, C. necator JMP134 2,4-dichlorophenol hydroxylase (YP_025383); ClpB S1, Defluvibacter lusatiensis S1 2,4-dichlorophenol hydroxylase (CAD60255); TfdB MH, Sphingobium herbicidovorans MH dichlorophenol hydroxylase (CAF32816); PheA EST1001, Pseudomonas sp. strain EST1001 phenol monooxygenase (AAC64901); OhpB V49, Rhodococcus sp. strain V49 3-(2-hydroxyphenyl) propionic acid monooxygenase (AAF81824); OrfL3 DFB63, Terrabacter sp. strain DBF63 2,4-dichlorophenol hydroxylase (BAB78767); HbpA HBP1, Pseudomonas azelaica HBP1 2-hydroxybiphenyl-3-monooxygenase (AAB57640); MhqA JMP134, C. necator JMP134 methylhydroquinone hydroxylase (YP_298872); MhqA NF100, Burkholderia sp. strain NF100 methylhydroquinone hydroxylase (BAE46529); DntB DNT, Burkholderia sp. strain DNT 4-methyl-5-nitrocatechol monooxygenase (ABC00744); DntB R34, Burkholderia cepacia R34 4-methyl-5-nitrocatechol monooxygenase (AAL50019); DxnD RW1, Sphingomonas sp. strain RW1 2,4-dihydroxybenzoate monooxygenase (CAA51370); ORF1 ATCC 39723, Sphingobium chlorophenolicum ATCC 39723 2,4-dihydroxybenzoate monooxygenase (AAM96655); CadA TQ07, P. putida TQ07 2,4-dihydroxybenzoate monooxygenase (AAL16082); PcpB ATCC 39723, S. chlorophenolicum ATCC 39723 pentachlorophenol-4-monooxygenase (AAF15368); PcpB MT1, Novosphingobium lentum MT1 pentachlorophenol-4-monooxygenase (CAC41015); ReutC6230 JMP134, C. necator JMP134 FAD-dependent monooxygenase (YP_293396); MobA KH122-3S, Comamonas testosteroni KH122-3S 3-hydroxybenzoate-4-hydroxylase (BAF34928); MhaA JMP134, C. necator JMP134 FAD-dependent monooxygenase (AAZ64667); MhaA U, P. putida U 3-hydroxyphenylacetate hydroxylase (AAY16572); ReutB4218 JMP134, C. necator JMP134 FAD-dependent monooxygenase (AAZ63571); NCgl1111 ATCC13032, Corynebacterium glutamicum ATCC13032 3-(3-hydroxyphenyl)propionate hydroxylase (NP_600384); ReutB5808 JMP134, C. necator JMP134 FAD-dependent monooxygenase (YP_299995); HppA PWD1, Rhodococcus globerulus PWD1 3-(3-hydroxyphenyl) propionate hydroxylase (AAB81312); MhpA K-12, Escherichia coli K-12 3-(3-hydroxyphenyl)propionate hydroxylase (NP_414881); MhpA JMP134, C. necator JMP134 3-(3-hydroxyphenyl)propionate hydroxylase (YP_294496); MhpA TA441, C. testosteroni TA441 3-(3-hydroxyphenyl)propionate hydroxylase (BAA82878); PobB CBS3, Pseudomonas sp. strain CBS3 4-hydroxybenzoate-3-monooxygenase (CAA52824); PobB GMI1000, Ralstonia solanacearum GMI1000 4-hydroxybenzoate-3-monooxygenase (NP_520363); PobB JMP134, C. necator JMP134 4-hydroxybenzoate-3-monooxygenase (YP_298206); PobA ADP1, Acinetobacter baylyi ADP1 4-hydroxybenzoate-3-monooxygenase (YP_046383); PobA B155, Rhizobium leguminosarum B155 4-hydroxybenzoate hydroxylase (AAA73519); PobA JMP134, C. necator JMP134 4-hydroxybenzoate hydroxylase (YP_299212); PobA E-37, Sagittula stellata E-37 4-hydroxybenzoate-3-monooxygenase (AAF65831); PobA2 ATCC 13525, Pseudomonas fluorescens p-hydroxybenzoate hydroxylase (AAA25834); PobA ATCC 13525, P. fluorescens 4-hydroxybenzoate-3-monooxygenase (CAA48483); PobA HR199, Pseudomonas sp. p-hydroxybenzoate hydroxylase (CAB43481); PobA ATCC 9043, Azotobacter chroococcum p-hydroxybenzoate hydroxylase (AAF03755); PobA KT2440, P. putida KT2440 4-hydroxybenzoate hydroxylase (AAN69138); ReutB5646 JMP134, FAD-dependent monooxygenase (YP_299835); MHPCO MA-1, Pseudomonas sp. MA-1 2-methyl-3-hydroxypyridine-5-carboxylic acid oxygenase (AAB60878); OnpA NyZ215, Alcaligenes sp. NyZ215 ortho-nitrophenol 2-monooxygenase (ABS81534); NahW AN10, Pseudomonas stutzeri AN10 salicylate-1-hydroxylase (AAD02157); NahG AN10, P. stutzeri AN10 salicylate-1-hydroxylase (AAD02146); NahG PpG7, P. putida PpG7 salicylate hydroxylase (YP_534831); SalA MT1, Pseudomonas sp. MT1 salicylate-1-hydroxylase (ABH07020); Sal S-1, P. putida S-1 salicylate hydroxylase (BAA61829); SalA ADP1, A. baylyi ADP1 salicylate-1-monooxygenase (YP_046111); MhbM1 JMP134, C. necator JMP134 3-hydroxybenzoate hydroxylase (YP_300048); MhbM2 JMP134, C. necator JMP134 3-hydroxybenzoate hydroxylase (YP_299992); MhbM M5a1, Klebsiella pneumoniae M5a1 3-hydroxybenzoate-6-hydroxylase (AAW63416); XlnD NCIMB 9867, Pseudomonas alcaligenes NCIMB9867 3-hydroxybenzoate-6-hydroxylase (AAG39455); ReutB3601 JMP134, C. necator JMP134 FAD-dependent monooxygenase (YP_297803); ORF7 ADP1, A. bayly ADP1 salicylate hydroxylase (YP_045698); NahG KT2440, P. putida KT2440 salicylate hydroxylase (NP_746074); ReutA2515 JMP134, C. necator JMP134 FAD-dependent monooxygenase (YP_296720); 6HNA3MO TN5, Pseudomonas fluorescens TN5 6-hydroxynicotinate-3-monooxygenase (E13001).

Two putative regulatory proteins are encoded in the pobAR-pcaLBGHQK gene cluster: (1) PobR, a XylS/AraC family regulator that might activate the expression of pobA gene in response to 4-hydroxybenzoate, as described in Azotobacter chroococcum ATCC 9043 (Quinn, 2001) and in P. putida WCS358 (Bertani, 2001); (2) pcaQ (C2 in Fig. 2), a LysR-type regulator homologous to the PcaQ regulator from Agrobacterium tumefaciens (Parke, 1996), that might control the expression of the pcaHGBL genes required to transform protocatechuate into β-ketoadipate (Fig. 1).

Peripheral pathways that channel phenylpropenoid and phenylpropanoid compounds to the β-ketoadipate pathway

Phenylpropenoid compounds are structural components of plant polymers, such as lignin and suberin, and constitute a common carbon source for plant-associated microorganisms. The ability to grow on phenylpropenoid compounds is widely distributed in bacteria. Among phenylpropenoid compounds, the largest group corresponds to hydroxycinnamates (i.e. ferulate, coumarate, caffeate and others). The hca genes, encompassing the hcaABCDEFG and hcaKR gene clusters, are responsible for the degradation of coumarate, caffeate and ferulate in A. baylyi ADP1 (Smith, 2003). The hcaG gene encodes a chlorogenate esterase that hydrolyzes the ester bond of chlorogenate, an abundant hydroxycinnamic compound, further producing quinate and caffeate (Fig. 1). Enzymes with a relatively broad substrate specificity – encoded by the hcaABC genes – carry out key steps in the dissimilation of coumarate, caffeate and ferulate (Fig. 1). HcaC, a hydroxycinnamoyl-CoA ligase, activates hydroxycinnamates to their thioester derivatives; HcaA, a bifunctional hydratase/lyase, converts the thioester derivative into an aldehyde intermediate; and HcaB, an aldehyde dehydrogenase, transforms the aldehyde to 4-hydroxybenzoate, protocatechuate and vanillate, respectively. The latter substrates are further degraded through the protocatechuate branch of the β-ketoadipate pathway (Fig. 1). The hcaRGKXABC gene cluster identified in C. necator (C2 in Fig. 2), has a relatively high identity to the Acinetobacter counterparts (Table 1). In addition to the hcaGABC genes, hcaK, a gene that encodes a putative transporter for hydroxycinnamate compounds, and hcaX, a gene that encodes a putative porin of unknown function, were also found in the hca cluster of C. necator. A putative repressor-encoding gene, hcaR, is located divergently from the hca operon. The hcaR gene is homologous to the A. baylyi ADP1 hcaR gene and related to the MarR-like family of transcriptional repressors (Table 1). By analogy with the hca gene cluster of A. baylyi ADP1 (Parke & Ornston, 2003), the inducers of the expression of hcaGKXABC genes in C. necator JMP134 may be the hydroxycinnamoyl-CoA thioesters.

It should be noted that ferulate allowed the growth of C. necator, although with a very low yield, but vanillate, an intermediate in the ferulate dissimilation pathway encoded by the hca genes, is not a growth substrate. This may be explained by the absence of demethylases in C. necator. In fact, none of the tested methylated compounds (vanillate, isovanillate, vanillin, vanillyl alcohol, isovanillyl alcohol, 2-, 3-, and 4-methoxybenzoates and 5-methoxysalicylate) allowed the growth of C. necator. In Pseudomonas and Acinetobacter species, a vanillate demethylase encoded by vanAB gene (Priefert, 1997; Segura, 1999) channels vanillate to protocatechuate. A genomic search for aromatic demethylase genes in C. necator renders only ORFs with a low identity with the vanAB genes of Pseudomonas and Acinetobacter strains. We hypothesize that C. necator JMP134 is unable to metabolize methoxylated aromatic compounds because it lacks the needed demethylase enzymes. The hca encoded functions convert ferulate to vanillate and acetyl-CoA. Acetyl-CoA formation would explain the weak growth and very low yield observed when ferulate is used as the growth substrate for C. necator. Phenylpropanoid compounds, i.e. saturated derivatives of hydroxycinnamates, such as 4-hydroxyphenylpropionate and 3,4-dihydroxyphenylpropionate serve as growth substrates for C. necator. In A. baylyi ADP1, it has been proposed that HcaD is a FAD-dependent acyl-CoA dehydrogenase which oxidizes the saturated propionyl-CoA side chain of the hydroxyphenylpropanoyl thioesters produced by HcaC (Smith, 2003), to form hydroxycinnamoyl-CoA thioesters; these are channeled by HcaA and HcaB gene products to the protocatechuate branch of the β-ketoadipate pathway (Fig. 1). In C. necator, a genomic search for homologues to the hcaD gene only renders ORFs with a very low identity with the A. baylyi ADP1 gene. It remains to be studied whether hydroxyphenylpropanoyl thioesters are transformed by a different acyl-CoA dehydrogenase in C. necator, to be further metabolized by the Hca gene products or are catabolized by a hca genes independent pathway.

Degradation of salicylate and 3-hydroxybenzoate: the gentisate pathway

In addition to 4-hydroxybenzoate, 2-hydroxybenzoate (salicylate) and 3-hydroxybenzoate also support the growth of C. necator JMP134. It has been well documented that, in Pseudomonas and Acinetobacter species, salicylate is oxidatively decarboxylated to produce catechol by salicylate-1-hydroxylase, a flavoprotein monooxygenase (Fig. 5) (You, 1991; Lee, 1996; Bosch, 1999; Jones, 2000), or by a three-component salicylate-1-hydroxylase in Sphingomonas species (Fig. 6) (Demaneche, 2004; Cho, 2005). In Burkholderia– a genus closely related to Cupriavidus– the conversion of salicylate into catechol has been also demonstrated (Hamzah & Al-Baharna, 1994); the hydroxylase has been purified (Ramsay, 1992) and the gene has been cloned, although not sequenced (Kim & Tu, 1989). The search in the genome of C. necator only showed two ORFs with low identity (c. a 30% aa identity) with salicylate 1-hydroxylase genes from gammaproteobacterial Pseudomonas and Acinetobacter strains. Furthermore, most salicylate-1-hydroxylases described so far are active with 4-chloro- and 5-chlorosalicylate which, in combination with a functional chlorocatechol ortho ring-cleavage pathway, such as that encoded in the plasmid pJP4 (see ‘Catabolic pathway for mono- and dichlorinated compounds: the tfd genes’), would allow degradation of chlorosalicylates. The fact that a salicylate 1-hydroxylase is absent in C. necator JMP134 is supported by the failure of the strain to grow on chlorosalicylates. An alternative route of salicylate degradation, via gentisate as an intermediate, is initiated by a salicylate-5-hydroxylase, as has been recently described in Ralstonia sp. U2 (Zhou, 2002). In contrast to salicylate-1-hydroxylases, salicylate-5-hydroxylase is a multicomponent enzyme consisting of an oxygenase, comprising the NagG and H subunits and an electron transport chain, comprising NagAa ferredoxin reductase, and NagAb ferredoxin. A putative salicylate-5-hydroxylase gene cluster comprising the genes hybRBCDA was identified (Table 2 and Fig. 2, C2) in the genome of strain JMP134. The putative LysR-type transcriptional regulator encoding hybR gene, and genes coding for the large (hybB) and small (hybC) subunits of the oxygenase as well as the ferredoxin encoding gene (hybD), show a significant similarity with the nagGHAb genes of Ralstonia sp. U2 and the hybBCD genes of Pseudomonas aeruginosa JB2 (Table 2). The hybBCD genes encode the second confirmed example of salicylate-5-hydroxylase activity carried out by a multicomponent oxygenase (Hickey, 2001). In contrast, the HybA gene product in strain JMP134 does not show any similarity to the NagAa protein from Ralstonia sp. U2 or to the HybA protein from P. aeruginosa JB2, which indicates a different evolutionary origin. However, the HybA gene product from strain JMP134 shows a similarity to MocF, the ferredoxin reductase of rhizopine demethylase in Rhizobium leguminosarum bv. viciae strain 1a (Bahar, 1998) and to AndAa, the ferredoxin reductase of a three-component anthranilate 1,2-dioxygenase in Burkholderia cepacia DBO1 (Chang, 2003). A further biochemical topic yet to be investigated is whether HybA and NagAa gene products are interchangeable in supplying the ferredoxin reductase function to the NagGHAb and HybBCD proteins, respectively.

Figure 6

Dendrogram showing the relatedness of Rieske nonheme iron oxygenase α subunits. The dendrogram was obtained by the neighbor-joining method using mega 4.0 based on sequence alignments calculated by clustal w using the default options. Sequences of deduced proteins encoded in the genome of Cupriavidus necator JMP134 are highlighted black. The sequences and their accession numbers are as follows: BenA JMP134, C. necator JMP134 benzoate-1,2-dioxygenase α subunit (AAZ63755); BenA ADP1, Acinetobacter baylyi ADP1 benzoate-1,2-dioxygenase α subunit (YP_046122); BenA KT2440, Pseudomonas putida KT2440 benzoate dioxygenase, α subunit (NP_745305); XylX mt-2, P. putida mt-2 toluate-1,2-dioxygenase α subunit (NP_542871); BenA RHA1, Rhodococcus sp. strain RHA1 benzoate-1,2-dioxygenase ISP α subunit (BAB70698); BenA ATCC 39116, Streptomyces setonii ATCC 39116 benzoate dioxygenase α subunit (AAN76670); CbdA TH2, Burkholderia sp. strain TH2 2-halobenzoate dioxygenase large subunit (BAB21584); CbeA NK8, Burkholderia sp. strain NK8 chlorobenzoate-1,2-dioxygenase α subunit (BAB21463); BenA TH2, Burkholderia sp. strain TH2 catechol-1,2-dioxygenases (BAC16780); XylX P2, Sphingomonas sp. strain P2 salicylate-1-hydroxylases (BAC65430); XylX F199, Novosphingobium aromaticivorans F199 toluate/benzoate dioxygenase large subunit (AAD04005); AntA ADP1, A. baylyi ADP1 anthranilate dioxygenase large subunit (CAG69424); AntA CA10, Pseudomonas resinovorans CA10 anthranilate-1,2-dioxygenase large subunit (BAC41526); TftA AC1100, Burkholderia cepacia AC1100 2,4,5-trichlorophenoxyacetic acid oxygenase (AAB39767); CadA HW13, Bradyrhizobium sp. strain HW13 2,4-D oxygenase large subunit (BAB78521); AtdA3 YAA, Acinetobacter sp. strain YAA aniline dioxygenase α-subunit (BAA13012); TadA1 AD9, Delftia tsuruhatensis AD9 salicylate-5-hydroxylase (AAX47241); TdnA1 7N, Delftia acidovorans 7N aniline dioxygenase large subunit (BAD61049); NdoB NCIB9816, P. putida NCIB9816 naphthalene-1,2-dioxygenase large subunit (NP_863072); HcaE K-12, Escherichia coli K-12 phenylpropionate dioxygenase large subunit (AAG57651); TodC1 F1, P. putida Fl phenylpropionate dioxygenase large subunit (ZP_00898191); BphA LB400, Burkholderia xenovorans LB400 biphenyl-2,3-dioxygenase α subunit (YP_556409); HybB JB2, Pseudomonas aeruginosa JB2 2-hydroxybenzoate-5-hydroxylase α subunit (AAC69484); NdsB NDS-2, Pigmentiphaga sp. strain NDS-2 salicylate-5-hydroxylase large subunit (BAC53590); NagG U2, Ralstonia sp. strain U2 salicylate-5-hydroxylase large oxygenase component (AAD12607); HybB JMP134, C. necator JMP134 salicylate-5-hydroxylase (YP_298911); ReutB3776 JMP134, C. necator JMP134 Rieske nonheme iron oxygenase (YP_297978); AndAc DBO1, B. cepacia DBO1 anthranilate dioxygenase large subunit (AAO83639); TerZa T7, D. tsuruhatensis T7 terephthalate-1,2-dioxygenase oxygenase large subunit (BAC15591); BphA1c F199, N. aromaticivorans F199 salicylate-5-hydroxylase large oxygenase component (NP_049213); AhdA1c P2, Sphingomonas sp. strain P2 salicylate-1-hydroxylase (BAC65426); OhbB JB2, P. aeruginosa JB2 ortho-halobenzoate-1,2-dioxygenase α (AAD20006); PhnA1b A5, Cycloclasticus sp. strain A5 salicylate-5-hydroxylase (BAC81540); BphA1d F199, N. aromaticivorans F199 aromatic oxygenase large subunit (NP_049206); AhdA1d P2, Sphingomonas sp. strain P2 salicylate-5-hydroxylase (BAC65433); ReutB5781 JMP134, C. necator JMP134 Rieske nonheme iron oxygenase (YP_299968); IphA2 YZW-D, C. testosteronii YZW-D isophthalate dioxygenase (AAX18934); Pht3 NMH102-2, P. putida NMH102-2 phthalate-4,5-dioxygenase (Q05183); OphA2 DBO1, B. cepacia DBO1 phthalate dioxygenase (AAD03558); CbaA BR60, C. testosteroni BR60 3-chlorobenzoate-3,4-dioxygenase oxygenase subunit (Q44256); ReutC6324 JMP134, C. necator JMP134 Rieske nonheme iron oxygenase (YP_293484); ReutB4795 JMP134, C. necator JMP134 Rieske nonheme iron oxygenase (YP_298987); PobA ROB310, Pseudomonas pseudoalcaligenes POB310 phenoxybenzoate dioxygenase (CAA55400); MnbA JS46, Comamonas sp. strain JS46 3-nitrobenzoate oxygenase (AAV33660); ReutA1473 JMP134, C. necator JMP134 Rieske nonheme iron oxygenase (YP_295687); ReutB3570 JMP134, C. necator JMP134 Rieske nonheme iron oxygenase (YP_297772); ReutB5789 JMP134, C. necator JMP134 Rieske nonheme iron oxygenase (AAZ65132); VanA NL15-2K, Streptomyces sp. strain NL15-2 K oxygenase α subunit (BAF33363); ReutD6483 JMP134, C. necator JMP134 Rieske nonheme iron oxygenase (YP_025403); VanA ADP1, A. baylyi ADP1 vanillate demethylase (AAC27107); VanA HR199, Pseudomonas sp. strain HR199 vanillate-O-demethylase (CAA72287); VanA BR6020, C. testosteroni BR6020 vanillate demethylase (AAW33713); DdmC DI6, Stenotrophomonas maltophilia DI-6 dicamba demethylase (AAV53699); IvaA BR6020, C. testosteroni BR6020 isovanillate demethylase (AAW33716); ReutB4340 JMP134, C. necator JMP134 Rieske nonheme iron oxygenase (YP_298537); TsaM T-2, C. testosteroni T-2 toluenesulfonate methyl-monooxygenase (AAC44804); TsaM2 T-2, C. testosteroni T-2 toluenesulfonate methyl-monooxygenase (AAK37996).

View this table:
Table 2

Genes encoding gentisate and homogentisate pathways and peripheral reactions

GenePosition (bp)No. aaRelated gene products
NameFunction/descriptionOrganism% Id (aa)Accession no.References
hybRC2 1420186-1419284300nagRLysR-like regulator proteinRalstonia sp. U2 pWWU257 (300)AAG13636Jones (2003)
hybB C2 1420295-1421548417nagGSalicylate-5-hydroxylase large oxygenase componentRalstonia sp. U2 pWWU278 (407)AAD12607Zhou (2002)
hybC C2 1421553-1422023156nagHSalicylate-5-hydroxylase small oxygenase componentRalstonia sp. U2 pWWU256 (160)AAD12608Zhou (2002)
hybDC2 1422036-1422350104hybDSalicylate-5-hydroxylase ferredoxinPseudomonas aeruginosa JB270 (94)AAC69486Hickey (2001)
hybAC2 1422382-1423641419mocFPutative ferredoxin reductaseRhizobium leguminosarum 1a pSyma42 (412)AAC31188Bahar (2000)
mhbD2C2 2625612-2626658348mhbDGentisate-1,2-dioxygenaseKlebsiella pneumoniae M5a160 (338)AAW63413Liu (2005)
mhbH2C2 2626758-2627456232mhbHFumarylpyruvate hydrolaseKlebsiella pneumoniae M5a170 (230)AAW63414Liu (2005)
mhbI2C2 2627478-2628119213mhbIMaleylpyruvate isomeraseKlebsiella pneumoniae M5a156 (211)AAW63415Liu (2005)
mhbM2C2 2628152-2629363403mhbM3-Hydroxybenzoate-6-hydroxylaseKlebsiella pneumoniae M5a162 (389)AAW63416Liu (2005)
mhbTC2 2699543-2698200447mhbTPutative 3-hydroxybenzoate transporterKlebsiella pneumoniae M5a151 (436)AAW63412Liu (2005)
mhbM1 C2 2700858-2699644404mhbM3-Hydroxybenzoate-6-hydroxylaseKlebsiella pneumoniae M5a161 (391)AAW63416Liu (2005)
mhbI1C2 2701526-2700885213mhbIMaleylpyruvate isomeraseKlebsiella pneumoniae M5a159 (211)AAW63415Liu (2005)
mhbH1C2 2702254-2701556232mhbHFumarylpyruvate hydrolaseKlebsiella pneumoniae M5a173 (230)AAW63414Liu (2005)
mhbD1C2 2703395-2702349348mhbDGentisate-1,2-dioxygenaseKlebsiella pneumoniae M5a159 (338)AAW63413Liu (2005)
mhbRC2 2703536-2704477313mhbRLysR-like regulatory proteinKlebsiella pneumoniae M5a140 (298)AAW63411Liu (2005)
mhaA C2 2086980-2088665561mhaA3-Hydroxyphenylacetate hydroxylase large componentPseudomonas putida U43 (527)AAY16572Arias-Barrau (2005)
hpdC2 1173069-1174148359hpd4-Hydroxyphenylpyruvate dioxygenasePseudomonas putida U79 (358)AAO12525Arias-Barrau (2004)
aroPC2 1174289-1175674461aroPAromatic amino acid transport proteinEscherichia coli K-1270 (451)P15993Honore & Cole (1990)
tyrBC2 1175795-1176994399tyrBTyrosine aminotransferaseEscherichia coli K-1257 (397)AAA24703Kuramitsu (1985)
phhAC1 3714366-3715316316phhAPhenylalanine-4-hydroxylaseChromobacterium violaceum69 (272)P30967Onishi (1991)
phhBC1 3715434-3715736100phhBPterin-4-α-carbinolamine dehydrataseRalstonia solanacearum GMI100071 (94)Q8XU38Salanoubat (2002)
hmgAC2 554469-555788439hmgAHomogentisate-1,2-dioxygenasePseudomonas putida U61 (427)AAO12527Arias-Barrau (2004)
hmgB1C2 555867-557135422hmgBFumarylacetoacetate hydrolasePseudomonas putida U49 (433)AAO12528Arias-Barrau (2004)
hmgB2C1 361952-362653233h16_A0361Putative fumarylacetoacetate hydrolaseCupriavidus necator H1694 (233)CAJ91512Pohlmann (2006)
hmgC C1 362658-363305215h16_A0362Maleylacetoacetate isomeraseCupriavidus necator H1684 (215)CAJ91513Pohlmann (2006)
  • * Genes for those cases where genetic or biochemical evidence for function is available are underlined.

A second gene cluster, ReutB3775-B3778, with homology to salicylate-5-hydroxylase components has been found in C. necator (Fig. 6). However, its identity with nag genes of Ralstonia sp. U2 or with the hyb genes found in P. aeruginosa JB2 (Hickey, 2001), is much lower (c. 40–45% aa identity) than that of the hyb genes cluster found in strain JMP134, which ranges between a 56% and a 78% aa identity (Table 2). The ReutB3775-B3778 gene cluster in strain JMP134 also shows a significant similarity with the ant gene cluster, which encodes anthranilate 1,2-dioxygenase of B. cepacia DBO1 (c. 35–45% aa identity) and with the three-component salicylate-1-hydroxylases (c. 35–50% aa identity), described in Sphingobium sp. P2 (Pinyakong, 2003). The actual role of this gene cluster in strain JMP134 remains to be elucidated.

In Proteobacteria, the following processes for the dissimilation of 3-hydroxybenzoate have been described: (1) in Pseudomonas alcaligenes (Poh & Bayly, 1980), Delftia acidovorans (Harpel & Lipscomb, 1990), B. cepacia (Wang, 1987), Klebsiella pneumoniae (Jones & Cooper, 1990; Suarez, 1995) and Salmonella typhimurium (Goetz & Harmuth, 1992), 3-hydroxybenzoate is degraded through gentisate by the activity of a 3-hydroxybenzoate-6-hydroxylase (Fig. 7); (2) in Comamonas testosteroni (Michalover, 1973; Hiromoto, 2006) and Bacillus sp. (Mashetty, 1996), the same compound is degraded through protocatechuate by a 3-hydroxybenzoate-4-hydroxylase. No ORFs closely related to the 3-hydroxybenzoate-4-hydroxylase gene (mobA) from C. testosteroni KH122-3S were found in the genome of strain JMP134 (Fig. 5). On the other hand, in the genome of this strain, two ORFs (mhbM1 and mhbM2) were identified with a significant (85% aa) identity with the 3-hydroxybenzoate-6-hydroxylase gene sequences from P. alcaligenes NCIB 9867 (xlnD) (Gao, 2005) and from K. pneumoniae M5a1 (mhbM) (Liu, 2005) (Table 2). The genes that encode the 3-hydroxybenzoate-6-hydroxylase in strain JMP134 (mhbM1 and mhbM2) cluster in the same group as those encoding the salicylate-1-hydroxylases, but in different branches in the dendrogram of FAD-dependent monooxygenases (Fig. 5). Both ORFs are located in putative gene clusters that encode enzymes involved in gentisate degradation (Fig. 2, C2, Table 2), thus supporting the possibility that C. necator JMP134 degrades 3-hydroxybenzoate through the gentisate pathway, as was suggested in an early report for C. necator 335 (Johnson & Stanier, 1971).

Figure 7

The gentisate, the homogentisate pathway, and their peripheral reactions.

The gentisate pathway (Fig. 7) is initiated by a gentisate-1,2-dioxygenase, which cleaves the aromatic ring between the carboxyl and the vicinal hydroxyl group to form maleylpyruvate. The latter compound can be converted into central metabolites of the Krebs cycle either by cleavage to pyruvate and maleate – performed by a maleylpyruvate hydrolase in a glutathione-independent way (Crawford & Frick, 1977; Bayly, 1980) – or by isomerization to fumarylpyruvate, via a glutathione-dependent maleylpyruvate isomerase (Zhou, 2001) (Fig. 7). In strain JMP134, both mhb gene clusters (Fig. 2, Table 2) contain putative mhbI and mhbH genes that are homologous to genes encoding maleylpyruvate isomerase (nagL) and fumarylpyruvate hydrolase (nagK) in Ralstonia sp. U2 (Zhou, 2001), respectively. This strongly suggests that gentisate is metabolized by a glutathione-dependent pathway in C. necator JMP134. The reason for the presence of two putative gene clusters encoding the enzymes required for the degradation of 3-hydroxybenzoate is not clear. The identity between homologous gene products from both clusters is very high (from a 78% aa identity in MhbI1/MhbI2 gene products to a 95% aa identity in MhbH1/MhbH2 gene products) and the gene order, mhbDHIM, is similar in both clusters (Fig. 2). However, the genomic context for each cluster is different. The first mhb gene cluster is located downstream of a LysR-type regulator gene, mhbR, and includes a mhbT gene that encodes a putative 3-hydroxybenzoate transporter (Fig. 2). The second cluster (Fig. 2) is associated at both ends with genes related to ATP-binding cassette (ABC)-type transporters, and is close to genes putatively encoding a FAD-dependent hydroxylase – moderately related to 3-hydroxyphenylpropionate hydroxylase – and an extradiol dioxygenase. It should be noted that two sets of isofunctional enzymes for the gentisate pathway, including 3-hydroxybenzoate-6-hydroxylases, have been described in P. alcaligenes NCIB 9867; one set is constitutively expressed, whereas the other set is strictly inducible by gentisate (Poh & Bayly, 1980). A similar situation may take place in strain JMP134. The mhbR gene is located divergently from the first mhb gene cluster and could be responsible for the induction of this or of both mhb gene clusters. The second mhb gene cluster is not associated to a regulatory gene and could be constitutively expressed, as has been shown in P. alcaligenes NCIB 9867 (Poh & Bayly, 1980).

The catabolic pathways for benzene, toluene, and (methyl)phenols

Conversion of benzene, toluene and (methyl)phenols into catechol

Cupriavidus necator JMP134 is able to grow on toluene, benzene, phenol, 2-, 3- and 4-methylphenols, 4-ethylphenol, 2,3- and 3,4-dimethylphenol. Some of these catabolic abilities have been described previously and are in agreement with our results (Pieper, 1995; Lang, 1996). Benzene, toluene, ethylbenzene and xylenes, commonly referred to as BTEX, are important nonoxygenated aromatic pollutants often found as mixtures at contaminated sites where fuels, solvents, or chemicals that are confirmed or suspected carcinogens, even at very low concentrations, have been spilled (Dean, 1985).

The utilization profile of methylphenols (2-, 3-, 4-methylphenol, 2,3- and 3,4-dimethylphenol) in C. necator JMP134 is similar to that of strains harboring a multicomponent phenol hydroxylase coupled with a catechol meta ring-cleavage pathway (see next section), as has been described for Pseudomonas sp. CF600 (Shingler, 1989). The (dimethyl)phenol hydroxylases catalyze the conversion into the corresponding catechols (Shingler, 1989). Multicomponent phenol hydroxylases belong to an evolutionary related family of soluble diiron hydroxylases, including enzymes involved in monooxygenation of inactive compounds such as methane and toluene, that lack an electron donating hydroxyl group (Leahy, 2003). The enzyme complexes consist of: an electron transport system comprising a reductase (and in some cases a ferredoxin); a catalytic effector protein containing neither organic cofactors nor metal ions, that is assumed to assemble an active oxygenase; a terminal hydroxylase with a (αβγ)2 quaternary structure; and a diiron center contained in each α-subunit (Leahy, 2003). Recently, these monooxygenases have been classified, according to their α-subunits, into four phylogenetic groups (Leahy, 2003): the soluble methane monooxygenases, the alkene monooxygenase of Rhodococcus corallinus B-276, the phenol hydroxylases and the four-component alkene/aromatic monooxygenases. Remarkably, two phenol hydroxylase encoding genes are found in the genome of C. necator JMP134. Genes that encode one phenol hydroxylase (phl1) (C1 in Fig. 3, Table 3), are associated with the catA2 gene, which encodes catechol-1,2-dioxygenase (see ‘The cat and ben genes’). This organization is highly similar to that of the mop genes operon of Acinetobacter calcoaceticus NCIB8250 (Ehrt, 1995). Therefore, it can be suggested that the catechol produced by the phl1 phenol hydroxylase gene cluster is metabolized through the ortho ring-cleavage pathway (Fig. 1), as has been described for phenol degradation in A. calcoaceticus NCIB8250. A second phenol hydroxylase gene cluster, phl2, is encoded by genes located downstream of the genes that encode a catechol meta ring-cleavage pathway (Table 3, C2 in Fig. 2), which also suggests a functional association. A growth rate-dependent expression of phenol assimilation pathways has been reported in C. necator JMP134 growing in continuous culture (Müller & Babel, 1996). At low growth rates, the ortho ring-cleavage pathway is almost exclusively expressed, but at high growth rates both ring-cleavage pathways are equally expressed. Induction of catechol-1,2-dioxygenase and catechol-2,3-dioxygenase is also detected in phenol-grown cells in batch cultures (Pieper, 1989; Kim & Harker, 1997); this indicates that C. necator JMP134 uses both phenol hydroxylases and catechol ring-cleavage pathways to grow on phenol, and perhaps on some methylphenols (Fig. 3). Analysis of the phl2 phenol hydroxylase showed the highest identities (83–96% aa, depending of the subunit, Table 3), with phenol hydroxylases of B. cepacia JS150 and Ralstonia pickettii PKO1, whose genes are associated with those of the meta ring-cleavage pathway. The phl1 phenol hydroxylase is highly identical to the corresponding enzyme of Wautersia numadzuensis TE26 (Table 3), and has lower identities (45–75% aa, depending on the subunit) with phenol hydroxylases of Ralstonia strains E2 (Hino, 1998) and KN1, (Nakamura, 2000), of the Comamonas strains TA441 (Arai, 1998) and R5 (Teramoto, 1999), and of the alkylphenol-degrading strains Pseudomonas sp. KL28 (Jeong, 2003) and P. putida MT4 (Takeo, 2006). Remarkably, the phl1 phenol hydroxylase of strain JMP134, and that of W. numadzuensis TE26, are the only ones whose genes are associated with a catechol-1,2-dioxygenase gene (note that the catechol-1,2-dioxygenase gene of strain TE26 is wrongly classified as a catechol-2,3-dioxygenase in the Genbank accession AB177762).

View this table:
Table 3

Genes encoding catabolic pathways for benzene, toluene, (methyl)phenols, 3-hydroxyphenylpropionate and 3-hydroxyanthranilate

Related gene products
GenePosition (bp)No. aaNameFunction/descriptionOrganism% Id (aa)Accession no.References
phlSC2 2502261-2501533242aphSNegative regulatory proteinComamonas testosteroni TA44155 (218)BAA89295Arai (1999a, b)
phlR2C2 2503952-2502258564aphRXylR/DmpR-type transcriptional regulatorComamonas testosteroni TA44156 (562)BAA34177Arai (1998)
tbcXC2 2504364-2505734456tbuXPutative membrane proteinRalstonia pickettii PKO178 (458)AAF03168Kahng (2000)
tbcRC2 2507565-2505787592tbuTTranscriptional activator TbuTRalstonia pickettii PKO188 (587)AAC44567Byrne & Olsen (1996)
tbcFC2 2508798-2507800332tbuCReductase subunitRalstonia pickettii PKO176 (334)AAS48552Fishman (2004)
tbcEC2 2509845-2508847332tbuA2β-Hydroxylase subunitRalstonia pickettii PKO187 (329)AAS48551Fishman (2004)
tbcDC2 2510175-2509858105tbuVEffector subunitRalstonia pickettii PKO195 (100)AAS48550Fishman (2004)
tbcCC2 2510530-2510195111tbuBFerredoxin subunitRalstonia pickettii PKO192 (111)AAS48549Fishman (2004)
tbcBC2 2510798-251053886tbuUγ-Hydroxylase subunitRalstonia pickettii PKO190 (86)AAS48548Fishman (2004)
tbcAC2 2512328-2510823501tbuA1α-Hydroxylase subunitRalstonia pickettii PKO193 (501)AAS48547Fishman (2004)
phlK2 C2 2512944-251315369crpAPhenol/cresol hydroxylase subunitRalstonia pickettii PKO183 (56)AAB67105Olsen (1997)
phlL2 C2 2513207-2514214335crpBPhenol/cresol hydroxylase subunitRalstonia pickettii PKO184 (334)AAB67106Olsen (1997)
phlM2 C2 2514253-251452289crpCPhenol/cresol hydroxylase subunitRalstonia pickettii PKO196 (89)AAB67107Olsen (1997)
phlN2C2 2514572-2516116514tbc1DTbc1D monooxygenaseBurkholderia cepacia JS15092 (514)AAG40791Kahng (2001)
phlO2C2 2516113-2516481122tbc1ETbc1E monooxygenaseBurkholderia cepacia JS15084 (66)AAG40792Kahng (2001)
phlP2C2 2516486-2517550354tbc1FTbc1F monooxygenaseBurkholderia cepacia JS15086 (354)AAG40793Kahng (2001)
phlQC2 2517567-2517932121cbzTFerredoxinPseudomonas putida GJ3170 (109)AAD05249Tropel (2002)
phlBC2 2517955-2518899314aphBCatechol-2,3-dioxygenaseComamonas testosteroni TA44183 (314)BAA34176Arai (2000)
phlXC2 2518948-2519397149orfYUnknownComamonas testosteroni TA44158 (143)BAA88499Arai (2000)
phlCC2 2519447-2520901484aphC2-Hydroxymuconic semialdehyde dehydrogenaseComamonas testosteroni TA44171 (484)BAA88501Arai (2000)
phlDC2 2520898-2521749283dmpD2-Hydroxymuconic semialdehyde hydrolasePseudomonas putida CF600 pVI15069 (272)CAA36993Nordlund & Shingler (1990)
phlEC2 2521778-2522560260aphE2-Hydroxypent-2,4-dienoate hydrataseComamonas testosteroni TA44166 (260)BAA88502Arai (2000)
phlHC2 2522586-2523377263aphH4-Oxalocrotonate decarboxylaseComamonas testosteroni TA44167 (261)BAA88505Arai (2000)
phlIC2 2523558-252374963aphI4-Oxalocrotonate isomeraseComamonas testosteroni TA44182 (63)BAA88507Arai (2000)
phlGC2 2652487-2651465340dmpG4-Hydroxy-2-oxovalerate aldolasePseudomonas putida CF600 pVI15084 (337)CAA43227Shingler (1992)
phlFC2 2653442-2652501313dmpFAcetaldehyde dehydrogenase (acylating)Pseudomonas putida CF600 pVI15085 (310)CAA43226Shingler (1992)
phlRC1 1834618-1832996540phtRPhenol hydroxylase regulator proteinWautersia numadzuensis TE2699 (442)BAD86546Kageyama (2005)
phlK1 C1 1834957-183521184phtAPhenol hydroxylase componentWautersia numadzuensis TE2691 (84)BAD86547Kageyama (2005)
phlL1 C1 1835259-1836281340phtBPhenol hydroxylase componentWautersia numadzuensis TE2693 (341)BAD86548Kageyama (2005)
phlM1 C1 1836298-183657090phtCPhenol hydroxylase componentWautersia numadzuensis TE2698 (90)BAD86549Kageyama (2005)
phlN1 C1 1836608-1838200530phtDPhenol hydroxylase componentWautersia numadzuensis TE2697 (530)BAD86550Kageyama (2005)
phlO1 C1 1838220-1838582120phtEPhenol hydroxylase componentWautersia numadzuensis TE2695 (120)BAD86551Kageyama (2005)
phlP1 C1 1838599-1839657352phtFPhenol hydroxylase componentWautersia numadzuensis TE2696 (352)BAD86552Kageyama (2005)
catA2C1 1839703-1840620305C12DCatechol-1,2-dioxygenaseWautersia numadzuensis TE2695 (305)BAD86553Kageyama (2005)
mhpRC1 299782-299000260mhpRmhp operon transcriptional activatorEscherichia coli K-1249 (253)CAA70746Ferrandez (1997)
mhpAC1 300051-301907618mhpA3-(3-Hydroxyphenyl)propionate hydroxylaseComamonas testosteroni TA44164 (572)BAA82878Arai (1999a, b)
mhpBC1 301910-302854314mhpB3-(2,3-Dihydroxyphenylpropionate)1,2-dioxygenaseEscherichia coli K-1255 (310)CAA70748Ferrandez (1997)
mhpDC1 302875-303663262mhpD2-Keto-4-pentenoate hydrataseComamonas testosteroni TA44162 (254)BAA82880Arai (1999a, b)
mhpTC1 303708-304979423mhpT3-Hydroxyphenylpropionic acid transporterEscherichia coli K-1249 (392)CAA66145Ferrandez (1997)
mhpCC1 305039-305908289mhpC2-Hydroxy-6-ketonone-2,4-dienedioic acid hydrolaseEscherichia coli K-1270 (282)CAA70749Ferrandez (1997)
haaRC2 2295021-2294074315nbaRRegulatory protein, LysR familyPseudomonas fluorescens KU-745 (312)BAC65313Muraki (2003)
haaEC2 2295167-2296630487nbaE2-Aminomuconate 6-semialdehyde dehydrogenasePseudomonas fluorescens KU-769 (486)BAC65304Muraki (2003)
haaHC2 2296637-2297461274nbaH2-Oxopent-4-dienoate hydratasePseudomonas fluorescens KU-765 (264)BAC65306Muraki (2003)
haaGC2 2297463-2298263266nbaG4-Oxalocrotonate decarboxylasePseudomonas fluorescens KU-766 (224)BAC65309Muraki (2003)
haaFC2 2298314-2298745143nbaF2-Aminomuconate deaminasePseudomonas fluorescens KU-772 (141)BAC65310Muraki (2003)
haaCC2 2298747-2299316189nbaC3-Hydroxyanthranilate 3,4-dioxygenasePseudomonas fluorescens KU-764 (170)BAC65311Muraki (2003)
haaDC2 2299313-2300326337nbaD2-Amino-3-carboxymuconate 6-semialdehyde decarboxylasePseudomonas fluorescens KU-767 (330)BAC65312Muraki (2003)
  • * Genes for those cases where genetic or biochemical evidence for function is available are underlined.

Multicomponent phenol hydroxylases display a significant oxidizing activity with trichloroethylene and have attracted considerable attention for their potential applications in bioremediation of this important pollutant. Also, the strain JMP134 exhibits significant trichloroethylene-oxidizing activity (Kim, 1996). It has been found that trichloroethylene degradation by phenol-degrading bacteria can be classified into three distinct kinetic groups: low-Ks (the half-saturation constant in Haldane's equation), moderate-Ks and high-Ks, which strictly correspond to three phylogenetic groups that are defined according to the phenol hydroxylase α-subunit sequence alignment (Futamata, 2001a). Consequently, both phenol hydroxylase α-subunits (PhlN) from C. necator JMP134 (Fig. 8a) fall into the group with a low Ks constant for trichloroethylene, which is considered the most useful group for an effective bioremediation of trichloroethylene-contaminated groundwater (Futamata, 2001b).

Figure 8

Dendrograms of α subunit multicomponent diiron monooxygenases. (a) (Methyl)phenol monooxygenases; (b) phenylacetyl-EoA oxygenases; and (c) toluene/benzene monooxygenases. Dendrograms were obtained by the neighbor-joining method using mega 4.0 based on sequence alignments calculated by clustal w using the default options. Sequences of deduced proteins encoded in the genome of Cupriavidus necator JMP134 are highlighted black. The sequences and their accession numbers are as follows: (A) LapN BH72, Azoarcus sp. BH72 phenol 2-monooxygenase p3 component (YP_933352); LapN KL28, Pseudomonas sp. strain KL28 phenol hydroxylase component (AAP92392); BupA4 MT4, Pseudomonas putida MT4 butylphenol hydroxylase component (BAC75404); PhlD H, P. putida H phenol hydroxylase (CAA56743); PhhN P35X, P. putida P35X phenol hydroxylase (CAA55663); DmpN CF600, Pseudomonas sp. strain CF600 phenol-2-monooxygenase (D37831); PhcN M1, Pseudomonas sp. M1 phenol hydroxylase subunit (AAY88747); PhN OX1, Pseudomonas stutzeri OX1 phenol hydroxylase component (AAO47358); MopN NCIB8250, Acinetobacter calcoaceticus NCIB8250 phenol hydroxylase component (CAA85383); MopN S13, A. radioresistens S13 phenol hydroxylase oxygenase component (AAM77905); DmpN II PM1, Methylibium petroleiphilum PM1 phenol hydroxylase α subunit (YP_001022494); PhyC KN1, Ralstonia sp. strain KN1 phenol hydroxylase component (BAA84121); PoxD E2, Ralstonia sp. strain E2 phenol hydroxylase component (AAC32455); DmpNI PM1, M. petroleiphilum PM1 phenol hydroxylase α subunit (YP_001021473); PhlN2 JMP134, C. necator JMP134 phenol hydroxylase (AAZ61067); PhtD TE26, Wautersia numadzuensis TE26 phenol hydroxylase component (BAD86550); PoxD BH72, Azoarcus sp. BH72 phenol hydroxylase (YP_933945); BtxD PHS1, Ralstonia sp. strain PHS1 oxygenase α subunit (ABG82169); TomA3 G4, Burkholderia cepacia G4 hydroxylase α subunit (AAL50373); AphN TA441, Comamonas testosteroni TA441 phenol hydroxylase component (BAA34172); PhcN R5, C. testosteroni R5 phenol hydroxylase subunit (BAA87871); PhkD KP23, Burkholderia kururiensis KP23 phenol hydroxylase subunit (BAB79282); Tbc1D JS150, Burkholderia cepacia JS150 monooxygenase (AAG40791); PhlN1 JMP134, C. necator JMP134 phenol hydroxylase (AAZ65028). (B) PaaG U, P. putida U phenylacetate-CoA oxygenase (AAC24334); PaaG2 Y2, Pseudomonas sp. strain Y2 phenylacetate-CoA oxygenase (CAE45110); PaaG Y2, Pseudomonas sp. strain Y2 phenylacetate-CoA oxygenase (CAD76929); PaaG ST, Pseudomonas fluorescens ST phenylacetate-CoA oxygenase (ABF82239); PaaA2 JMP134, C. necator JMP134 phenylacetate-CoA oxygenase (AAZ62566); PaaA GMI1000, Ralstonia solanacearum GMI1000 phenylacetate-CoA oxygenase (NP_522165); PaaA (PacE) KB740, Azoarcus evansii KB740 phenylacetate-CoA oxygenase (CAC10606); PaaA EbN1, Azoarcus sp. strain EbN1 phenylacetate-CoA oxygenase (YP_159026); PaaA CGA009, Rhodopseudomonas palustris CGA009 phenylacetate-CoA oxygenase (NP_949105); PaaA USDA110, Bradyrhizobium japonicum USDA110 phenylacetate-CoA oxygenase (NP_769531); PaaA JMP134 (Reut_B3735), C. necator JMP134 phenylacetate-CoA oxygenase (AAZ63093); PaaA2 EbN1, Azoarcus sp. strain EbN1 phenylacetate-CoA oxygenase (YP_160273); PaaA W, Escherichia coli W phenylacetate-CoA oxygenase (P76077); PaaA PAMU-1.2, Klebsiella sp. strain PAMU-1.2 phenylacetate-CoA oxygenase (BAE02686) (C) TmoA KR1, Pseudomonas mendocina KR1 toluene-4-monooxygenase (AAS66660); BtxP PHS1, Ralstonia sp. strain PHS1 monooxygenase α subunit (ABG82181); TbhA AA1, B. cepacia AA1 toluene-3-monooxygenase oxygenase subunit 1 (AAB58740); BmoA Jl104, P. aeruginosa Jl104 benzene monooxygenase oxygenase subunit (BAA11761); TbuA1 PM1, M. petroleiphilum PM1 toluene monooxygenase α subunit (YP_001020011); TouA OX1, Pseudomonas sp. strain OX1 toluene-o-xylene monooxygenase component (AAT40431); TbcA JMP134, C. necator JMP134 toluene/benzene hydroxylase (AAZ65024); Tbc2A JS150, B. cepacia JS150 monooxygenase (AAG40794); TbuA1 PKO1, Ralstonia pickettii PKO1 hydroxylase α subunit (AAS48547).

Another phenol hydroxylase gene cluster associated with a catechol-1,2-dioxygenase gene has been described in A. calcoaceticus NCIB8250 (Ehrt, 1995). However, the amino acid identities of the phenol hydroxylase multicomponent of C. necator JMP134 with the phenol hydroxylase of A. calcoaceticus are lower than the identities with the meta ring-cleavage-associated phenol hydroxylases indicated above. This suggests that the clustering of phenol hydroxylase operons with catechol-1,2-dioxygenase genes occurred independently and after the divergence of both phenol hydroxylases. In addition, the amino acid identity between the catechol-1,2-dioxygenases from both clusters is not particularly high (54%), as compared with the amino acid identities (c. 70%) of the CatA2 gene product and the catechol-1,2-dioxygenases that are not directly related to phenol hydroxylase gene clusters (Fig. 4).

Based on the overall sequence identity and gene order, it is reasonable to propose that PhlN1/PhlN2 (Fig. 8a), PhlL1/PhlL2, and PhlO1/PhlO2 gene products correspond to the α, β, and γ subunits of the putative (αβγ)2 hexameric nonheme diiron monooxygenases, respectively. Both putative α subunits have the highly conserved motif Asp–Glu–X–Arg–His, which occurs twice in the PhlN1/PhlN2 gene products at positions 148/139 and 243/234, respectively. These ligands form the dinuclear iron-binding site in the large subunits of this family of monooxygenases (Fox, 1988). In addition, the spacing of 94 aa between these motifs is conserved in both PhlN gene products.

PhlP1/PhlP2 gene products show identity with a very large group of iron–sulfur flavoproteins that transfer electrons from reduced pyridine nucleotides to a terminal electron acceptor via a flavin and [2Fe–S] center (Mason & Cammack, 1992). PhlM1/PhlM2 gene products are homologous to the small polypeptides that are believed to play a role in regulating monooxygenase activity, namely the effector proteins. The only member of this group of proteins that has been studied in detail is the DmpM protein of Pseudomonas sp. CF600 (Qian, 1997). DmpM protein binds to the DmpNLO hydroxylase, but does not participate directly in redox reactions. Rather, its role seems to be to increase the steady state turnover rates and the product yields from the phenol hydroxylase, possibly by controlling the entrance of substrate and exit of products (Qian, 1997). PhlM1/PhlM2 gene products from C. necator JMP134 possess the conserved amino acids Glu56/Glu54 and Gly59/Gly57, respectively, which were identified as conserved components of the functionally important helix 2 of the DmpM protein (Qian, 1997). However, Leu56 of the DmpM protein is replaced by Met58 in the PhlM1 gene product and by Thr56 in the PhlM2 gene product, which indicates that the conservation of this residue is not required.

The PhlK1/PhlK2 gene products are unique, because they share significant similarity only with a small group of polypeptides that are exclusively found as components of phenol hydroxylases, and not as components of other multicomponent diiron monooxygenases. The role of the PhlK1/PhlK2 gene products can be inferred from studies on the DmpK protein from Pseudomonas sp. CF600 (Powlowski, 1997). The DmpK protein binds to both DmpN and DmpL proteins and plays an essential role in the assembly of the active form of the oxygenase – possibly by posttranslational insertion of iron into the subunit – because the DmpK protein can catalyze in vitro reactivation of the inactive enzyme in the presence of iron.

A multicomponent monooxygenase that belongs to an alkene/aromatic monooxygenase subfamily and putatively encodes a toluene/benzene/xylene monooxygenase (tbc genes, C2 in Fig. 2) is located divergently from the phl2 gene cluster in C. necator. The analysis of the deduced protein products of this tbcABCDEF gene cluster revealed a strong similarity to the nearly identical tbc2 gene cluster of Burkholderia sp. JS150 and to the tbu gene cluster of R. pickettii PKO1 (76–95% aa, Table 3). The TbcA/TbuA1/Tbc2A gene products cluster in a separate branch in the dendrogram of α subunits of the toluene/benzene monooxygenases (Fig. 8c), and have been involved in the hydroxylation of inactivated aromatic compounds like benzene, toluene and o-xylene. Three other strains have a similar arrangement of phenol hydroxylase and toluene/benzene/xylene monooxygenase encoding genes: Burkholderia sp. JS150 (Kahng, 2001), R. pickettii PKO1 (Byrne, 1995), and Pseudomonas stutzeri OX1 (Bertoni, 1998). In the latter strain, as has been recently proposed, the coupling of the two enzymatic systems optimizes the use of nonhydroxylated aromatic molecules by the draining effect of the phenol hydroxylase on the product(s) of the oxidation catalyzed by toluene monooxygenase; this avoids phenol accumulation (Cafaro, 2004).

The toluene/benzene monooxygenase gene cluster of strain JMP134 has been cloned previously, partially sequenced, and expressed (Kim, 1996). The plasmid pYK3021, which contains the cloned toluene/benzene monooxygenase gene cluster, exhibited phenol hydroxylase activity. However, it is not possible to establish if this activity corresponds to the toluene/benzene monooxygenase [as has been shown for the toluene/o-xylene monooxygenase of P. stutzeri OX1 (Bertoni, 1998)] or to the presence of the phl2 gene cluster that encodes the meta ring-cleavage pathway associated phenol hydroxylase, in the same cloned 9.1-kb fragment. This is why the genes encoding the toluene/benzene/xylene monooxygenase in strain JMP134 were originally labeled as phl (GenBank accession AF065891) (Kim, 1996). Given the genomic information presented here and the high identity with the toluene/benzene/xylene monooxygenases from Burkholderia sp. JS150 and P. stutzeri OX1 (Table 3), we consider the phl denomination incorrect, and have changed it to tbc genes, leaving the former phl denomination for the phenol hydroxylase gene clusters of C. necator JMP134 (Table 3).

The enzymes of the toluene/benzene/xylene monooxygenase family consist of four dissociable components, three of which constitute a short electron transfer chain with an oxidoreductase, a ferredoxin, and a terminal hydroxylase. Based on the overall sequence and gene order similarity, it is reasonable to consider the TbcF gene product as the oxidoreductase, the TbcC gene product as the ferredoxin, and the TbcAEB gene products as the terminal hydroxylase, in the electron transfer chain of the Tbc monooxygenase (Table 3). The TbcF gene product from strain JMP134 is similar to the proteins that comprise the very large family of iron–sulfur flavoproteins; these function as oxidoreductases for most mono- and dioxygenase systems. Supporting this assumption is the presence, in the TbcF gene product, of the conserved Cys (Cys37, -42, -45 and -77) and Gly (Gly40 and -52) residues that are required for the coordination of the two iron atoms of the [2Fe–2S] cluster (Mason & Cammack, 1992). The TbcC gene product is highly homologous to the ferredoxin component of other toluene/benzene/xylene monooxygenases. This group of ferredoxins is related to a large family of Rieske-type ferredoxins that function as soluble electron carriers for a variety of bacterial oxygenases. The TbcC gene product contains the metal-binding motif Cys–X–His–X15–21–Cys–X2–His at positions 44–66, which is characteristic of all Rieske-type proteins (Mason & Cammack, 1992). A high degree of similarity with the components of the terminal hydroxylases of multicomponent monooxygenases suggests that the TbcAEB gene products correspond to the (αβγ)2 dimeric nonheme iron monooxygenase. In agreement with this, in the TbcA gene product, the putative α-subunit of the monooxygenase, there are two copies of the motif (Asp/Glu)–X26–30–Asp–Glu–X–Arg–His at positions 104–137 and 197–234; these are the ligands for the diiron center in the active sites of the enzymes of this family (Fox, 1994).

The TbcD gene product appears homologous to the small polypeptides that might play a role in regulating the monooxygenase activity. Among the toluene/benzene/xylene monooxygenases, the TmoD protein from Pseudomonas mendocina KR1 has been studied in detail (Pikus, 1996). The TmoD protein, which appears to be a substoichiometric constituent of the TmoAEB hydroxylase, can mildly stimulate the rate of toluene hydroxylation when added to purified hydroxylase. Moreover, the TbcD gene product from strain JMP134 has, at positions Glu67, Leu70 and Gly71, the amino acids identified as conserved components of the functionally important helix 2 of the DmpM protein, which provides a similar regulatory function for the phenol hydroxylase of Pseudomonas sp. CF600 (Qian, 1997). Therefore, it is reasonable to conclude that the TbcD gene product might function as a regulatory component of the Tbc monooxygenase complex.

The regulation of the expression of the putative phenol hydroxylases and the toluene/benzene/xylene monooxygenase of strain JMP134 seems to be complex. The ortho ring-cleavage pathway associated with the Phl1 phenol hydroxylase has a regulatory gene encoded divergently from the putative phl1 genes operon (C1 in Fig. 2). This is the classical organization in most multicomponent phenol hydroxylases. The product of this gene, PhlR1, belongs to the aromatic-responsive σ54-dependent family of regulators that includes the well-characterized DmpR transcriptional activator involved in the regulation of phenol degradation in Pseudomonas sp. CF600 (Shingler, 1993; Shingler & Moore, 1994). On the other hand, three putative regulatory genes are located next to the tbc genes cluster: tbcR, phlR2 and phlS (C2 in Fig. 2). The tbcR and phlR2 genes encode regulators that also belong to the aromatic-responsive σ54-dependent family. Both PhlR1 and PhlR2 gene products have the highest identity with the PoxR and AphR proteins, which are positive regulators of phenol degradative operons from W. numadzuensis TE26, and C. testosteroni TA441, respectively (Table 3). These regulators are responsive to phenol or alkylphenols, because members of this family exhibit broad effector-response profiles (Shingler & Moore, 1994). On the other hand, the TbcR gene product, putatively involved in the regulation of the tbc gene cluster, has a high amino acid identity with the TbuT protein (88%); TbuT is the activator of the toluene monooxygenase tbu genes operon of R. pickettii PKO1, which is activated not only by aromatic effectors as benzene, toluene or ethylbenzene, but also by trichloroethylene (Kahng, 2000). The PhlS gene product, the third putative regulator encoded in the region of the phl2 and tbc gene clusters in C. necator, has significant identity with regulators belonging to the GntR family of transcriptional repressors, like the aphS or phcR genes from C. testosteroni strains TA441 (Arai, 1999b) and R5 (Teramoto, 2001), respectively. In the absence of the genuine substrate, these regulators repressed the gratuitous expression of phenol-metabolizing enzymes. The presence of several regulators in C. necator, putatively involved in phenol degradation, suggests a complex regulatory system that comprises cross-regulation, regulatory cascades, competition for binding sites and regulatory hierarchy. Finally, between the tbcR and phlR2 genes, it was possible to identify the tbcX gene (Table 3), whose gene product is almost identical (99% in amino acid) with the TbuX protein from R. pickettii PKO1 (Kahng, 2000), an outer membrane protein that plays an important role in the catabolism of toluene.

The catechol meta ring-cleavage pathway

The biochemical route of the meta ring-cleavage pathway for the degradation of catechol produced by phenol hydroxylation is illustrated in Fig. 3. The putative functions of the phl genes encoded enzymes are shown in Table 3. The critical ring-opening step of the meta ring-cleavage pathway is typically catalyzed by type I catechol-2,3-dioxygenase enzymes, the so called vicinal oxygen chelate family enzymes (Vaillancourt, 2004), that usually contain nonheme Fe+2 at the active site. Inactivation during turnover of para-substituted catechols (Cerdan, 1995) and 3-chlorocatechol has been described for members of this family. The phlB gene-encoded enzyme shares the highest identity with catechol-2,3-dioxygenases of the subfamily 1.2.C of extradiol dioxygenases (Fig. 9a) in the classification system proposed by Eltis and Bolin (Eltis & Bolin, 1996). Two genes, named mpcI and mpcII– that encode enzymes with activity against catechol derivatives – had been cloned previously from C. necator JMP222, a pJP4 cured derivative of C. necator JMP134 (Kabisch & Fortnagel, 1990a, b). The Km values for catechol of these two enzymes are in the millimolar range; this is unexpected for catechol-2,3-dioxygenases and indicates that catechol is not the native substrate. None of these two mpc genes correspond to the phlB gene, the catechol-2,3-dioxygenase encoding gene involved in phenol degradation. The genome analysis established that the mpcI gene should be designated mhpB (Fig. 9b), given that the gene product does not belong to the vicinal oxygen chelate superfamily, but shows a high similarity to 2,3-dihydroxyphenylpropionate-1,2-dioxygenases (Bugg, 1993) (see ‘The 2,3-dihydroxyphenylpropionate meta ring-cleavage pathway), which makes it part of the type II extradiol dioxygenases that exhibit only poor activity with catechol (Spence, 1996).

Figure 9

Dendrograms of extradiol dioxygenases. (a) Subfamily I.2 type I extradiol dioxygenases; (b) type II extradiol dioxygenases; and (c) subfamilies I.3, I.4 and I.6 type I extradiol dioxygenases. Dendrograms were obtained by the neighbor-joining method using mega 4.0 based on sequence alignments calculated by clustal w using the default options. Sequences of deduced proteins encoded in the genome of Cupriavidus necator JMP134 are highlighted black. The sequences and their accession numbers are as follows: (A): PhnE1 RP007, Burkholderia sp. strain RP007 catechol-2,3-dioxygenase (AAF02426); Cdo AA1, Burkholderia cepacia AA1 catechol-2,3-dioxygenase (AAB88079); Cdo MT15, Pseudomonas putida MT15 catechol-2,3-dioxygenase (AAC43044); CbzE GJ31, P. putida GJ31 catechol-2,3-dioxygenase (AAD05250); CdoE JS765, Comamonas sp. strain JS765 catechol-2,3-dioxygenase (AAC79918); AphB TA441, Comamonas testosteroni TA441 catechol-2,3-dioxygenase (BAA34176); PhlB JMP134, C. necator JMP134 catechol-2,3-dioxygenase (YP_299876); TdnC UUC2, P. putida UCC2 3-methylcatechol-2,3-dioxygenase (CAA42452); TadC1 AD9, Delftia tsuruhatensis catechol-2,3-dioxygenase (AAX47246); TomB G4, Burkholderia vietnamiensis G4 catechol-2,3-dioxygenase (YP_001110005); BtxH PHS1, Ralstonia sp. strain PHS1 catechol-2,3-dioxygenase (ABG82173); NahH AN10, Pseudomonas stutzeri AN10 catechol-2,3-dioxygenase (AAD02148); XylE mt-2, P. putida mt-2 metapyrocatechase (CAA24490); NahH G7, P. putida G7 catechol-2,3-dioxygenase (YP_534833); DmpB CF600, Pseudomonas sp. strain CF600 catechol-2,3-dioxygenase (P17262); PhlH H, P. putida H catechol-2,3-dioxygenase (CAA56747); AtdB YAA, Acinetobacter sp. strain YAA catechol-2,3-dioxygenase (BAA23555); PheG KN1, Ralstonia sp. strain KN1 catechol-2,3-dioxygenase (BAA84125); CmpE HV3, Sphingomonas sp. strain HV3 catechol-2,3-dioxygenase (CAB06613); XylE B1, Beijerinckia sp. strain B1 catechol-2,3-dioxygenase (B57264); TdnC2 UCC2, P. putida UCC2 catechol-2,3- dioxygenase (BAB62050); LapB KL28, Pseudomonas sp. strain KL28 catechol-2,3-dioxygenase (AAP92388); BupB MT4, P. putida MT4 catechol-2,3-dioxygenase (BAC75400) (B) HpaD W, Escherichia coli W homoprotocatechuate dioxygenase (CAA86042); AmnA AP-3, Pseudomonas sp. strain AP-3 2-aminophenol-1,6-dioxygenase (BAB03532); AmnB AP-3, Pseudomonas sp. strain AP-3 2-aminophenol-1,6-dioxygenase (BAB03531); MhpB ML2, P. putida ML2 2,3-dihydroxyphenylpropionate-1,2-dioxygenase (AAG09232); MhpB W3110, E. coli W3110 2,3-dihydroxyphenylpropionate-1,2-dioxygenase (BAA13053); MhpB JMP134, C. necator JMP134 catechol-2,3-dioxygenase (YP_294497); OhpD V49, Rhodococcus sp. strain V49 3-(2,3-dihydroxyphenyl) propionic acid dioxygenase (AAF81826); ORFK1 DFB63, Terrabacter sp. strain DBF63 meta ring-cleavage enzyme (BAB91220); HppB PWd1, Rhodococcus globerulus PWD1 2,3-dihydroxyphenylpropionate-1,2-dioxygenase (AAB81314); EdoD I1, Rhodococcus sp. strain I1 catechol-2,3-dioxygenase (CAA06875); MhpB TA441, C. testosteroni TA441 3-(2,3-dihydroxyphenyl)propionate-1,2-dioxygenase (BAA82879); GalA KT2440, P. putida KT2440 gallate dioxygenase (NP_744666); PcmA 12B, Arthrobacter keyseri 12B protocatechuate-4,5-dioxygenase (AAK16524); PmdB BR6020, C. testosteroni BR6020 protocatechuate-4,5-dioxygenase β subunit (AAK73573); FldU LB216, Sphingomonas sp. strain LB126 protocatechuate dioxygenase β subunit (CAB87561); LigB SYK-6, Sphingomonas paucimobilis SYK-6 protocatechuate-4,5-dioxygenase β subunit (BAB88743); PhnC RP007, Burkholderia sp. strain RP007 catechol-2,3-dioxygenase (AAD09870); FlnD1 DBF63, Terrabacter sp. strain DBF63 2′-carboxy-2,3-dihydroxybiphenyl 1,2-dioxygenase (BAE45095); CarBb GTIN11, Sphingomonas sp. strain GTIN11 2-aminobiphenyl-2,3-diol-1,2-dioxygenase (AAL37978); CarBb CA10, Pseudomonas resinovorans CA10 anthranilate-1,2-dioxygenase (BAC41547); ReutB4784 JMP134, C. necator JMP134 extradiol dioxygenase (AAZ64132); LigZ SYK-6, S. paucimobilis SYK-6 biphenyl meta ring-cleavage enzyme (BAA31862); ReutB5775 JMP134, C. necator JMP134 extradiol dioxygenase (YP_299962); DesZ SYK-6, S. paucimobilis SYK-6 3-O-methylgallate-3,4-dioxygenase (BAC79261). (C): BphC KF715, P. putida KF715 2,3-dihydroxybiphenyl dioxygenase (AAA25756); BphC LB400, Burkholderia xenovorans strain LB400 2,3-dihydroxybiphenyl-1,2-dioxygenase (YP_556403); IpbC JR1, Pseudomonas sp. strain JR1 3-isopropylcatechol-2,3-dioxygenase (AAB36672); BphCI P6, Rhodococcus globerulus P6 2,3-dihydroxybiphenyl-1,2-dioxygenase I (CAA53297); TodE F1, P. putida F1 biphenyl-2,3-diol-1,2-dioxygenase (ZP_00898196); BphC RHA1, Rhodococcus sp. strain RHA1 2,3-dihydroxybiphenyl dioxygenase (BAA06872); ReutC6234 JMP134, C. necator JMP134 extradiol dioxygenase (YP_293400); ReutA1133 JMP134, C. necator JMP134 extradiol dioxygenase (YP_295355); MhqB JMP134, C. necator JMP134 extradiol dioxygenase (YP_298870); MhqB NF100, Burkholderia sp. strain NF100 extradiol dioxygenase (BAE46530); BphCII BN6, Sphingomonas xenophaga BN6 2,3-dihydroxybiphenyl dioxygenase (AAB58815); ORFI JB2, P. aeruginosa JB2 catechol-2,3-dioxygenase (AAC69491); BphC4 RHA1, Rhodococcus sp. strain RHA1 2,3-dihydroxybiphenyl-1,2-dioxygenase (BAA98135); BphC A401, Pseudomonas stutzeri A401 catechol-2,3-dioxygenase (BAA20161); BphC KC37, Pseudomonas mendocina KC37 catechol-2,3-dioxygenase (BAA20162); DntD DNT, B. cepacia DNT trihydroxytoluene oxygenase (AAD12738); DntD R34, B. cepacia R34 2,4,5-trihydroxytoluene oxygenase (AAF89669); Edi2 YK2, Rhodococcus sp. strain YK2 catechol-2,3-dioxygenase (BAC00798); BphC3 RHA1, Rhodococcus sp. strain RHA1 catechol-2,3-dioxygenase (YP_703930); ReutB5807 JMP134, C. necator JMP134 extradiol dioxygenase (YP_299994); DbfB RW1, Sphingomonas sp. strain RW1 catechol-2,3-dioxygenase (CAA51364); PcbC DJ-12, Pseudomonas sp. strain DJ-12 extradiol dioxygenase (BAA07956); HbpC HBP1, Pseudomonas azelaica HBP1 2,3-dihydroxybiphenyl-1,2-dioxygenase (AAB57639); NsaC BN6, S. xenophaga BN6 1,2-dihydroxynaphthalene dioxygenase (AAB06725); BphC F199, Novosphingobium aromaticivorans F199 dihydroxy naphthalene/biphenyl dioxygenase (NP_049210); EtbC RHA1, Rhodococcus sp. strain RHA1 2,3-dihydroxybiphenyl-1,2-dioxygenase (ABH00328); EdoA NCIMB13064, Rhodococcus rhodochrous NCIMB13064 catechol-2,3-dioxygenase (AAC18907).

The mpcII (ReutB4677 mhpB) gene – together with two other extradiol dioxygenase encoding genes found in the genome of C. necator JMP134 (ReutA1133, and ReutC6234, Fig. 9c) – is related to the bphCII gene of Sphingomonas xenophaga BN6 and to the bphC2 and bphC3 genes of Rhodococcus globerulus P6, which encode a new dimeric type of extradiol dioxygenase with 2,3-dihydroxybiphenyl dioxygenase activity (Heiss, 1997). The important feature of this group of enzymes is their small subunit molecular weight; only half as much as other types of extradiol dioxygenases. Phylogenetic analyzes indicated that the ancestral type I extradiol dioxygenase was, like the small size subunit enzymes, a one-domain enzyme, and that two-domain enzymes arose from a single duplication event (Eltis & Bolin, 1996). Unfortunately, no physiological function has been assigned to the BphCII gene product in the naphthalenesulfonate-degrading strain BN6 that clarify the role of this mpcII-like class of extradiol dioxygenases (Fig. 9c). In R. globerulus P6, the BphC2 gene product was constitutively expressed, and supposedly supported the degradation of PCBs (McKay, 2003).

A new gene, mhqB, related to mpcII-like dioxygenases was recently described in Burkholderia sp. NF100 and putatively considered to encode a extradiol dioxygenase involved in methylhydroquinone catabolism. Such protein showed an extradiol ring-cleavage activity toward 3-methylcatechol and a lower activity with catechol (Tago, 2005), suggesting possible physiological substrates of these enzymes in C. necator JMP134. Several mutagenesis attempts to obtain catechol-2,3-dioxygenase mutants of C. necator JMP134, which should show impaired growth on methylphenols, have been unsuccessful (D. Pieper, unpublished data); this suggests that if the phlB gene is inactivated, redundant functions supply the catechol-2,3-dioxygenase activity. The mpcII-like group of extradiol dioxygenases is the most likely candidate to undertake this function, as these enzymes usually exhibit some activity against 3-methylcatechol and very poor activity with catechol (Asturias & Timmis, 1993; McKay, 2003).

In the meta ring-cleavage pathway operon of strain JMP134 (C2 in Fig. 2), the phlB gene is preceded by the phlQ gene that encodes a putative ferredoxin with a high identity with the cbzT gene product from P. putida GJ31; this strain has a chlorocatechol-2,3-dioxygenase, the CbzE protein, which is exceptionally resistant to inactivation by 3-chlorocatechol (Kaschabek, 1998). These small auxiliary ferredoxin proteins – whose genes are frequently encoded adjacently to the catechol-2,3-dioxygenase genes in the meta-pathway operons – have a reactivating function, as has also been shown for the XylT protein in toluene catabolism, for the NahT protein in naphthalene catabolism and for the PhhQ and DmpQ proteins in methylphenol catabolism (Hugo, 2000). Similarly, the CbzT protein is able to reactivate the CbzE protein in vitro, through reduction of the iron atom, when the enzyme had been fully inactivated by 4-methylcatechol (Tropel, 2002). Loss of the activity of the DmpQ or XylT proteins, respectively, results in strains that are unable to grow on compounds that are catabolized through p-substituted methylcatechols (i.e. 4-methyl and 3,4-dimethylcatechol) (Polissi & Harayama, 1993; Powlowski & Shingler, 1994). However, these strains can still grow on compounds that are catabolized through catechol or 3-methylcatechol. Therefore, it can be proposed that the ability of strain JMP134 to metabolize 4-methylphenol and 3,4-dimethylphenol might depend on the regeneration of an active catechol-2,3-dioxygenase through the activity of the PhlQ gene product.

An ORF of unknown function, phlX, which encodes a relatively hydrophobic protein, is located downstream of the phlB gene. Similar genes have been found in other gene clusters encoding the meta ring-cleavage pathway, such as the phnX gene of Burkholderia sp. RP007, the orfY gene of C. testosteroni TA441, the cbzX gene of P. putida GJ31 and the nahX gene of plasmid NAH7 from P. putida G7 (Grimm & Harwood, 1999; Laurie & Lloyd-Jones, 1999; Mars, 1999; Arai, 2000). The role of these phlX-like genes has not been determined, but an orfY mutant of C. testosteroni TA441 grew poorly on phenol and accumulated the yellow 2-hydroxymuconic semialdehyde intermediate in the medium; this suggests that it could be involved in additional catabolism of 2-hydroxymuconic semialdehyde (Arai, 2000). The next two genes in the meta ring-cleavage pathway operon, phlC and phlD, encode 2-hydroxymuconic semialdehyde dehydrogenase (HMSD) and hydrolase (HMSH) and show a high identity with homologous gene products of the meta ring-cleavage pathway operon in Pseudomonas sp. CF600 (Table 3). Both enzymes use ring-cleavage products as substrates: 2-hydroxymuconic semialdehyde from catechol, 5-methyl-2-hydroxymuconic semialdehyde from 4-methylcatechol and 2-hydroxy-6-oxo-2,4-heptadienoate from 3-methylcatechol. However, 2-hydroxymuconic semialdehyde is only a poor substrate for HMSH, and it is preferentially degraded via the oxalocrotonate branch of the meta ring-cleavage pathway (Harayama, 1987). Because the ring-cleavage product of 3-methylcatechol is a ketone, rather than an aldehyde, it cannot be further oxidized by the HMSD and must be metabolized via the hydrolytic route (Fig. 3). Correspondingly, it has been shown that mutants of the HMSH encoding gene (dmpD) in Pseudomonas sp. CF600 still grew on phenol or 4-methylphenol but failed to grow on phenols that are channeled through 3-methylsubstituted catechols (Powlowski & Shingler, 1994). On the other hand, Pseudomonas sp. CF600 with a deletion in the HMSD gene (dmpC) or in either of the genes encoding the other two enzymes of the 4-oxalocrotonate branch (the dmpI and dmpH genes) resulted in strains that grew on 2-methyl-, 3-methyl- and 3,4-dimethylphenol, but not on phenol or 4-methylphenol. These results indicate that despite the potential use of either branch for the metabolism of the ring-cleavage products of catechol and 4-methylcatechol, these compounds are preferentially metabolized by the HMSD rather than by the hydrolase of the meta ring-cleavage pathway (Powlowski & Shingler, 1994). The same situation may take place with the different phenols that are growth substrates for C. necator. The last three genes identified in the meta ring-cleavage pathway operon in strain JMP134 are phlE, phlH and phlI; these encode: (1) 4-oxalocrotonate isomerase (4OI), which catalyzes the isomerization of the enol form of 4-oxalocrotonate to its keto-form; (2) 4-oxalocrotonate decarboxylase (4OD), which catalyzes the formation of 2-oxopent-4-dienoate, the common intermediate of the hydrolytic branch and the 4-oxoalocrotonate branch of the meta ring-cleavage pathway, and (3) 2-oxopent-4-dienoate hydratase (Fig. 3). All these genes in strain JMP134 show the highest identities with the homologous gene products of C. testosteroni TA441 (Arai, 2000). Gene order in the phl genes of the meta ring-cleavage pathway in C. necator JMP134 is clearly different from any other reported phl gene cluster. Moreover, it is the only example in which genes encoding the 4-hydroxy-2-ketovalerate aldolase (HOA, the phlG gene) and the aldehyde dehydrogenase (acylating, the phlF gene) – that catalyze the final steps of the meta ring-cleavage pathway to generate the end-products pyruvate and acetyl-CoA, respectively (Powlowski, 1993) – are separated from the rest of the other meta ring-cleavage pathway genes (C2 in Fig. 2). These genes show a high identity with the xylQ and xylK genes, respectively, from the pWW0 plasmid of P. putida, where they are encoded with the rest of the xyl genes (Assinder & Williams, 1990; Aemprapa & Williams, 1998).

The methylcatechol ortho ring-cleavage pathway

The degradation of methyl-substituted catechols as intermediates in the degradation of methylaromatics usually proceeds through a catechol meta ring-cleavage pathway. The metabolism of methylcatechols via the ortho ring-cleavage pathway results in the formation of methyl-substituted 4-carboxymethylbut-2-en-4-olides (methylmuconolactones) as dead-end products (Catelani, 1971; Knackmuss, 1976). In the transformation of 4-methylcatechol, 4-methylmuconolactone (4-ML) is formed, which cannot be processed by enzymes of the β-ketoadipate pathway as no proton is available to be abstracted by the muconolactone isomerase. In C. necator JMP134, however, a different ortho ring-cleavage pathway for the degradation of 4-methylcatechol has been described (Pieper, 1985) (Fig. 3). A key enzyme of this new pathway in C. necator JMP134 (Pieper, 1985, 1990) was initially characterized as a 4-ML methylisomerase capable of converting 4-ML to 3-methylmuconolactone (3-ML). This enzyme's function may be to compensate for the initial ‘incorrect’ cycloisomerization of 3-methylmuconate. 3-ML is further metabolized via 4-methyl-β-ketoadipate, and hence, probably, by analogous reactions to those of the classical β-ketoadipate pathway (Fig. 3).

A 3-kb mml gene cluster, harboring the gene that encodes 4-ML methylisomerase, was cloned and sequenced (Erb, 1998), and additional genes were identified (C1 in Fig. 2, Table 2). The first gene in the mml cluster, mmlH, encodes a putative transporter protein for 4-ML and exhibits a sequence homology to other members of the major superfamily of transmembrane facilitators, showing the common structural motif of 12 transmembrane-spanning α-helical segments, and the key amino acid motif that is characteristic of this superfamily (Erb, 1998). The second gene, mmlI, encodes the 4-ML methylisomerase and, given the novelty of the reaction, no sequence homologies were found. Finally, the mmlJ gene encodes a methylmuconolactone isomerase (Prucha, 1997) with significant identity to the muconolactone isomerases of Pseudomonas and Acinetobacter strains (Houghton, 1995) and of strain JMP134, which is involved in the degradation of catechol via the β-ketoadipate pathway. The methylmuconolactone isomerase encoded by the mmlJ gene is supposed to transform 3-ML produced from 4ML, by 4-ML methylisomerase, to 4-methyl-β-ketoadipate enol-lactone. Further metabolism of this intermediate and the possible formation of 4-methyl-β-ketoadipate are far from being understood.

By genomic analysis, it was possible to identify the genetic context of the mml genes in strain JMP134. Two genes, mmlFG, putatively encoding both subunits of a β-ketoadipate CoA transferase were found upstream to the mmlH gene (C1 Fig. 2, Table 1). Both genes show a high identity with the homologous genes (pcaIJ and catIJ genes) in the Pseudomonas and Acinetobacter strains (Shanley, 1994; Gobel, 2002), suggesting a possible role in the further metabolism of 4-methyl-β-ketoadipate enol-lactone. A putative LysR-type regulator encoding gene (mmlR) was found directly upstream of mmlFG, which indicates that mmlFG and mmlHIJ genes are part of one transcriptional unit (C2 in Fig. 2. Table 1). It should be noted that no gene that putatively encodes an isoenzyme of β-ketoadipate enol-lactone hydrolase was found in the mml gene cluster. This point supports the idea that 4-methyl-β-ketoadipate enol-lactone is not further metabolized through a classical β-ketoadipate pathway. Recently, a gene cluster was found on the megaplasmid pHG1 of C. necator H16 (Schwartz, 2003) with the same order of the mml genes in strain JMP134, and with amino acid identity levels of 80–90% (Table 1). This mml gene cluster also includes the genes that putatively encode the β-ketoadipate-CoA transferase and the LysR-type regulator.

Degradation of C6-C2 and C6-C3 compounds

The phenylacetyl-CoA ring-cleavage pathway

Although phenylacetate is a common source of carbon and energy for a wide variety of microorganisms, knowledge on the bacterial catabolism of this natural aromatic compound is still fragmentary, and details on the enzymatic mechanisms and the nature of intermediates are scarce. The general pathway for aerobic phenylacetate metabolism has initially been characterized in Gammaproteobacteria, Escherichia coli (Ferrandez, 1998), P. putida (Olivera, 1998) and the betaproteobacterium Azoarcus evansii (Mohamed Mel, 2002; Rost, 2002) (Fig. 10). This pathway does not follow the conventional route for the aerobic degradation of aromatics. In E. coli, there are 14 paa genes that encode for phenylacetate degradation, organized in three transcriptional units; two of them, paaZ and paaABCDEFGHIJK, encode the catabolic genes; the third, paaXY, contains the paaX regulatory gene (Ferrandez, 1998). In a study of paa mutants of E. coli K12, a phenylacetate catabolic pathway has been proposed (Ismail, 2003). The initial step of the pathway involves the activation of phenylacetate into phenylacetyl-CoA by a phenylacetate-coenzyme A ligase, encoded by the paaK gene (Ferrandez, 1998). The respective enzymes have also been identified in P. putida (Olivera, 1998) and A. evansii (El-Said Mohamed, 2000). The phenylacetyl-CoA is attacked by a ring-oxygenase/reductase (the PaaABCDE gene products), generating a hydroxylated and reduced derivative of phenylacetyl-CoA, which is not reoxidized to a dihydroxylated aromatic intermediate as in other known aromatic pathways (Fig. 10). It has been proposed that this nonaromatic intermediate CoA ester is further metabolized in a complex reaction sequence comprising enoyl-CoA isomerization/hydration, nonoxygenolytic ring opening and dehydrogenation, which is catalyzed by the PaaG and PaaZ gene products. The resulting aliphatic CoA dicarboxylate compound is further catabolized by a β-oxidation-like pathway via β-ketoadipyl-CoA into succinyl-CoA and acetyl-CoA, which appears to be catalyzed by the PaaF, PaaJ and PaaH gene products (Ismail, 2003).

Figure 10

The phenylacetate, the benzoate, and the anthranilate pathways proceeding through aryl-CoA intermediates.

A search in the genome of strain JMP134 showed 19 genes putatively involved in phenylacetate catabolism (Table 4), organized in three clusters (C1 and C2 in Fig. 2). Two genes were found, paaK1 and paaK2, which putatively encode phenylacetate CoA-ligase, both showing over a 70% aa identity with the PaaK gene product of A. evansii KB740 (Table 4). The amino acid sequence identity between both PaaK gene products is also over 70%, but the genetic context is completely different (Fig. 2). The analysis of the sequence of both paaK gene products revealed the presence of three conserved motifs for AMP and substrate binding in acyl-adenylate-forming enzymes (Ferrandez, 1998). The motif I comprises residues 97/96SSGTTGKPTV106/105 in the PaaK1/PaaK2 gene products, matching the AMP-binding site consensus sequence T(SG)-S(G)-G-(ST)-T(SE)-G(S)-X-P(M)-K-G(LAF) (predominant aa are underlined and alternatives are in parentheses) (Ferrandez, 1998). The sequences 239/238DIYGLSE245/244 and 305/304YRTRD309/308 in the PaaK1/PaaK2 gene products from C. necator JMP134, which are conserved in all putative or bona fide phenylacetate CoA-ligases, correspond to motifs II and III, respectively, and they may contribute to the substrate-binding sites (Ferrandez, 1998). It should be noted that two functional, almost identical copies of genes that encode phenylacetate-CoA ligases, each one located in a different genetic context, have also been reported in the styrene-degrading Pseudomonas sp. strain Y2 (Alonso, 2003); however, the physiological meaning of the existence of two phenylacetate-CoA ligases in this strain is not clear. Two gene clusters, paaA1B1C1D1E1 and paaA2B2C2D2E2, that putatively encode ring-oxygenase/reductase multicomponent proteins are also found in C. necator JMP134 (Table 4, and Fig. 2). The amino acid identity between the gene products of both gene clusters range from 45% to 65%. However, the PaaA1 and PaaA2 subunits of the multicomponent oxygenase map far apart in the corresponding dendrogram (Fig. 8b). Both gene clusters show the highest identities with different homologous clusters (Table 4); paaA1B1C1D1E1 is closer to the paaGHIJK gene cluster of Pseudomonas sp. strain Y2 (Alonso, 2003), whereas the paaA2B2C2D2E2 is closer to the paaABCDE (pacEFGHI) gene cluster of A. evansii KB740 (Mohamed Mel, 2002; Rost, 2002). It has been proposed that the paaABCDE genes encode a five-subunit oxygenase enzyme complex, using phenylacetyl-CoA as substrate (Ferrandez, 1998; Diaz, 2001; Mohamed Mel, 2002; Ismail, 2003; Fernández, 2006). PaaACD may function as a terminal oxygenase, with PaaA as the large α subunit containing the dinuclear iron-binding site (Ferrandez, 1998). The small protein PaaB may be the dissociable activator protein required for an optimal turnover of the oxygenase component (Ferrandez, 1998). Finally, the similarity of the PaaE protein to class IA-like reductases – members of the ferredoxin-NADP+ reductase family – indicate that it could function as a reductase delivering electrons from NAD(P)H to the terminal PaaACD oxygenase (Ferrandez, 1998). The phenylacetyl-CoA oxygenase constitutes the first reported multicomponent oxygenase acting on a CoA-activated aromatic compound (Fernández, 2006). Interestingly, although all bacterial multicomponent oxygenases described so far are monooxygenases, the product of the reaction catalyzed by the phenylacetyl-CoA oxygenase is a dihydrodiol, and therefore this enzyme could be a hydroxylating dioxygenase rather than a monooxygenase (Fernández, 2006). In C. necator JMP134, the presence of two gene clusters putatively encoding phenylacetyl-CoA multicomponent oxygenases has been reported. This is the first reported case for double dosage in these genes and its role is unclear. On the other hand, only one copy of the paaZ gene that putatively encodes the proposed ring-opening enzyme is found in the genome of strain JMP134 (C2 in Fig. 2). The PaaZ gene product from C. necator JMP134 shows only a 23% aa identity with the E. coli PaaZ. The E. coli protein has an N-terminal region (residues 1–527) that contains all the conserved motifs that characterize the aldehyde dehydrogenase superfamily, and a C-terminal domain with some similarity to enoyl-CoA hydratases (Diaz, 2001). Therefore, it has been proposed that the E. coli PaaZ is a fused, bifunctional protein with the enoyl-CoA hydratase-like C-terminal domain involved in the ring-cleavage of the phenylacetate intermediate, because enoyl-CoA hydratases have been linked with the ring-cleavage in the anaerobic benzoate degradation pathway (Diaz, 2001). The PaaZ gene product of C. necator JMP134 is a shorter polypeptide (554 aa) than the E. coli PaaZ protein (681 aa), and it lacks the enoyl-CoA hydratase-like C-terminal domain, which suggests that this paaZ gene is in a prefusion state. Close relatives of the C. necator JMP134 PaaZ gene product are the protein PaaZ (PacL) of A. evansii KB740 (Mohamed Mel, 2002; Rost, 2002) and the protein PaaN2 of Pseudomonas sp. Y2 (Alonso, 2003). In these bacteria, the ring-cleavage could not be undertaken by PaaZ, because the enoyl-CoA hydratase-like domain is also absent; therefore, PaaZ may participate in the conversion of the aldehyde, produced by the ring opening, into a carboxylic acid, as has been recently proposed in E. coli (Ismail, 2003). Given the similarity of the PaaG protein with some members of the enoyl-CoA hydratase/isomerase family, it has been proposed that, in E. coli, the ring opening may be preceded by a reversible PaaG-catalyzed Δ3, Δ2 isomerization of double bonds and/or by the addition of water in the putative cis-dihydrodiol derivative of phenylacetyl-CoA (Ismail, 2003). In addition, given that C–C-cleaving enoyl-CoA hydratases have been described, it has been proposed that PaaG may play a role in C–C cleavage (Ismail, 2003). Therefore, it is interesting to speculate that in C. necator JMP134 and A. evansii KB740 strains that possess an ‘aldehyde dehydrogenase only’ version of PaaZ, PaaG could be directly involved in the ring opening of the putative cis-dihydrodiol derivative of phenylacetyl-CoA (Fig. 10). The paaG gene of C. necator JMP134 is located close to the paaF gene, another putative enoyl-CoA hydratase encoding gene, paaI, that encodes a protein of unknown function and the paaK2 gene, that encodes a putative phenylacetyl-CoA ligase (Fig. 2). The amino acid identity with the E. coli paaG gene product is moderate (58%), and similar to the identity showed with the P. putida U paaB gene product (Table 4). In the mutational analysis of paa genes, performed on E. coli, in addition to paaZ and paaG, three additional genes proved to be essential for the utilization of phenylacetate as the carbon source: paaF, paaH and paaJ (Ismail, 2003), all of them putatively involved in the final steps in phenylacetate degradation (Fig. 10). The PaaH protein of E. coli is similar to 3-hydroxyacyl-CoA dehydrogenases, which suggests that it catalyzes the dehydrogenation of 3-hydroxyadipyl-CoA to produce β-ketoadipyl-CoA. On the other hand, the PaaF protein has a sequence similarity to proteins of the enoyl-CoA hydratase (isomerase) family (crotonase family), which may have cis-Δ3-trans-Δ2-enoyl-CoA isomerase activity, in addition to the enoyl-CoA hydratase activity (Ismail, 2003). Supposedly, in the absence of the PaaF and PaaH proteins, the catabolism of phenylacetate ends at the level of 3-hydroxyadipyl-CoA. Homologous genes to paaF and paaH are found in the genome of strain JMP134. The paaF gene is located close to the paaG gene and the paaH gene is found just next to the second copy of the paaG gene, paaG2 (Table 4). In E. coli, the β-ketoadipyl-CoA intermediate produced by the action of the PaaH protein could be thiolytically cleaved by the PaaJ/PaaE protein (which would be similar to the β-ketoadipyl-CoA thiolases involved in the lower part of the β-ketoadipate pathway) in order to produce acetyl-CoA and succinyl-CoA (Ismail, 2003; Nogales, 2007). It should be emphasized that no gene homologous to paaJ/paaE is located in the paa gene clusters of C. necator JMP134 (Fig. 2). Therefore, it is reasonable to assume that for the phenylacetate catabolism in this strain, the enzyme that should be encoded by the paaJ/paaE gene can be recruited from other catabolic pathways such as the β-ketoadipate pathway; specifically, the pcaF gene encoding a β-ketoadipyl-CoA thiolase (Fig. 10).

View this table:
Table 4

Genes encoding pathways for phenylacetate, benzoate, and anthranilate proceeding through aryl-CoA intermediates

Related gene products
GenePosition (bp)No. aaNameFunction/ descriptionOrganism% Id (aa)Accession no.References
padC2 716863-715364499styDPhenylacetaldehyde dehydrogenasePseudomonas sp. Y253 (500)CAA04003Velasco (1998)
aauAC2 718286-71777171aauAAromatic amine dehydrogenaseAlcaligenes faecalis57 (177)AF302652Chistoserdov (2001)
aauDC2 718893-718306195aauEAromatic amine dehydrogenaseAlcaligenes faecalis65 (191)AF302652Chistoserdov (2001)
aauEC2 719491-718928187aauEAromatic amine dehydrogenaseAlcaligenes faecalis38 (168)AF302652Chistoserdov (2001)
aauBC2 720693-719488401aauBAromatic amine dehydrogenaseAlcaligenes faecalis42 (335)AF302652Chistoserdov (2001)
paaXC2 328688-327765307paaX2Putative transcriptional repressorPseudomonas sp. Y238 (306)CAE45100Bartolome-Martin (2004)
paaA1 C2 328817-329800327paaGPutative ring-oxidation complex protein 1Pseudomonas sp. Y274 (324)CAD76929Velasco (1998)
paaB1 C2 329856-33013793paaHPutative ring-oxidation complex protein 2Pseudomonas sp. Y262 (93)CAD76932Velasco (1998)
paaC1 C2 330164-330916250paaIPutative ring-oxidation complex protein 3Pseudomonas sp. Y262 (246)CAD76933Velasco (1998)
paaD1 C2 330922-331437171paaJ2Putative ring-oxidation complex protein 4Pseudomonas sp. Y255 (162)CAE45113Bartolome-Martin (2004)
paaE1 C2 331456-332532358paaKPutative ring-oxidation complex protein 5Pseudomonas sp. Y258 (356)CAD76937Velasco (1998)
paaZC2 332597-334261554paaN2Putative ring-opening enzymePseudomonas sp. Y258 (547)CAD76913Velasco (1998)
paaK1C2 334348-335694448paaKAerobic phenylacetate-CoA ligaseAzoarcus evansii KB74076 (439)AAF26285El-Said Mohamed (2000)
paaPC2 335803-336117104paaP2Putative membrane proteinPseudomonas sp. Y248 (95)CAE45115Bartolome-Martin (2004)
paaLC2 336114-337778554paaL2Putative phenylacetic acid permeasePseudomonas sp. Y279 (520)CAE45116Bartolome-Martin (2004)
paaA2C1 3519904-3520908334pacEPutative ring oxidation complex protein 1Azoarcus evansii KB74074 (332)CAC10606Rost (2002)
paaB2C1 3520931-352121895pacFRelated to aerobic phenylacetate degradationAzoarcus evansii KB74077 (95)CAC10607Rost (2002)
paaC2C1 3521231-3522073280pacGPutative ring oxidation complex protein 2Azoarcus evansii KB74054 (262)CAC10608Rost (2002)
paaD2C1 3522087-3522629180pacHPutative ring oxidation complex protein 3Azoarcus evansii KB74055 (167)CAC10609Rost (2002)
paaE2C1 3522664-3523752362pacIFerredoxinAzoarcus evansii KB74056 (362)CAC10610Rost (2002)
paaRC1 3524766-3525401211paaRTranscriptional regulator tetR familyAzoarcus sp. EbN142 (209)CAI08130Rabus (2005)
paaFC1 3314249-3313473258paaFPutative enoyl-CoA hydratase proteinAzoarcus sp. EbN169 (252)CAI09380Rabus (2005)
paaGC1 3318495-3319319274paaBEnoyl-CoA hydratase IIPseudomonas putida U58 (258)AAC24330Olivera (1998)
paaIC1 3319366-3319812148paaD2Phenylacetic acid degradation proteinPseudomonas sp. Y250 (138)CAE45105Bartolome-Martin (2004)
paaK2C1 3319895-3321199434paaKAerobic phenylacetate-CoA ligaseAzoarcus evansii KB74071 (430)AAF26285El-Said Mohamed (2000)
paaG2C1 1097624-1098403259paaG2Putative enoyl-CoA hydratase proteinRalstonia solanacearum GMI100072 (259)CAD15716Salanoubat (2002)
paaHC1 1098446-1099969507paaH2Putative 3-hydroxybutyryl-CoA dehydrogenaseRalstonia solanacearum GMI100071 (501)CAD15715Salanoubat (2002)
boxAC1 1418515-1417262417boxABenzoyl-CoA oxygenase component AAzoarcus evansii KB74061 (422)AAN39377Zaar (2004)
boxBC1 1420015-1418591474boxBBenzoyl-CoA oxygenase component BAzoarcus evansii KB74071 (472)AAN39376Zaar (2004)
boxCC1 1421727-1420045560boxCEnoyl-CoA-hydratase/isomeraseAzoarcus evansii KB74070 (556)AAN39375Zaar (2004)
bzdRC1 1422765-1421836309bzdRRegulatory proteinAzoarcus sp. CIB53 (276)AAQ08805Lopez Barragan (2004)
bzdAC1 1423044-1424705553bzdABenzoate-CoA ligaseAzoarcus sp. CIB62 (511)AAQ08820Lopez Barragan (2004)
bzdXC1 1424719-1425534271ORF2Lactonase of the 3,6 lactone of 3-hydroxyadipyl-CoAAzoarcus evansii KB74051 (235)AAN39366Gescher (2002)
abmEC1 2404061-2403651136orf5Putative translation regulator proteinAzoarcus evansii KB74059 (132)AAL02075Schuhle (2001)
abmXC1 2404556-2404107149orf1ThioesteraseAzoarcus evansii KB74035 (138)AAN39365Gescher (2002)
abmGC1 2406199-2404553548orf72-Aminobenzoate-CoA ligaseAzoarcus evansii KB74058 (546)AAL02069Schuhle (2001)
abmDC1 2407427-2406258389orf4Putative acyl-CoA dehydrogenaseAzoarcus evansii KB74071 (382)AAL02066Schuhle (2001)
abmCC1 2408312-2407461283orf3Putative enoyl-CoA hydratase/isomeraseAzoarcus evansii KB74070 (277)AAL02065Schuhle (2001)
abmBC1 2409177-2408374267orf2Putative β-hydroxyacyl-CoA dehydrogenaseAzoarcus evansii KB74063 (260)AAL02072Schuhle (2001)
abmBC1 2409177-2408374267orf2Putative β-hydroxyacyl-CoA dehydrogenaseAzoarcus evansii KB74063 (260)AAL02072Schuhle (2001)
abmAC1 2411546-2409177789orf12-Amninobenzoyl-CoA monooxygenase/reductaseAzoarcus evansii KB74066 (783)AAL02071Schuhle (2001)
kynAC1 877146-878030294kynATryptophan-2,3-dioxygenaseRalstonia metallidurans CH3488 (299)EAN53667Kurnasov (2003)
kynUC1 878043-879299418kynUKynureninaseRalstonia metallidurans CH3483 (418)EAN53666Kurnasov (2003)
kynBC1 879332-879991219kynBKynurenine formamidase C-terminal fragmentRalstonia metallidurans CH3472 (129)EAN53665Kurnasov (2003)
KynBKynurenine formamidase N-terminal fragmentRalstonia metallidurans CH3482 (73)EAN53664Kurnasov (2003)
  • * Genes for those cases where genetic or biochemical evidence for function is available are underlined

The regulation of the paa gene cluster in E. coli is controlled by two elements: (1) PaaX, a transcriptional repressor which contains a stretch of 25 residues with similarity to the HTH motif of transcriptional regulators of the GntR family, but constitutes a different family; and (2) phenylacetyl-CoA, which specifically inhibits the binding of PaaX to the repression-binding sites in the promoter sequences of the paa cluster (Ferrandez, 2000). A PaaX-binding sequence is located divergently to the paaA1B1C1D1E1ZK1PL gene cluster in C. necator JMP134, which suggests that this paa gene cluster is regulated in a similar way to that of E. coli. It should be noted that downstream of the paaA2B2C2D2E2 genes in strain JMP134, a putative TetR-family transcriptional regulator gene (paaR) is encoded, whose gene product shows a 44% aa identity with ORF3 of the phenylacetate catabolism gene cluster of A. evansii KB740 (Rost, 2002); this indicates that this putative gene could also be involved in the regulation of this paa gene cluster.

In C. necator JMP134, the degradation of several aromatic compounds can be assumed to proceed through the phenylacetyl-CoA pathway. The genes that encode a periplasmic aromatic amine dehydrogenase (aau genes) were identified (Table 4; Fig. 2). These could be responsible for the catabolism of phenylethylamine into phenylacetaldehyde, which could then be transformed into phenylacetate by a phenylacetaldehyde dehydrogenase (Pad) (Fig. 10) (Chistoserdov, 2001). The degradation of phenylalkanoates that contain an even number of carbon atoms as phenylbutyrate and phenylhexanoate, is assumed to be accomplished by a β-oxidation complex which catalyzes the formation of phenylacetyl-CoA, as has been shown in P. putida (Olivera, 2001), but the presence and identity of these genes in C. necator JMP134 could not be determined. On the other hand, the metabolism of phenylpyruvate in Azospirillum brasilense (Somers, 2005) and Saccharomyces cerevisiae (Vuralhan, 2003) has been proposed to occur through decarboxylation into phenylacetate. In C. necator JMP134, however, putative genes for phenylpyruvate decarboxylase were not identified, which suggests the existence of a different pathway or a different kind of decarboxylase-encoding gene.

The homogentisate ring-cleavage pathway

The homogentisate ring-cleavage pathway is widespread in eukaryotic and prokaryotic cells, because it is the central route for phenylalanine and tyrosine catabolism. Phenylalanine is converted into tyrosine by a pterin- and metal-dependent phenylalanine hydroxylase (PhhA) with an auxiliary carbinolamine dehydratase (PhhB) that is responsible for the regeneration of the pterin cofactor. A tyrosine aminotransferase (TyrB) transforms tyrosine into 4-hydroxyphenylpyruvate, which is further converted into homogentisate by a 4-hydroxyphenylpyruvate dioxygenase (Hpd) (Fig. 7). Homologous genes for the phenylalanine and tyrosine peripheral pathways are present in the genome of strain JMP134 (Table 2). The phhA and phhB genes are clustered (C1 in Fig. 2), and show a 46% and a 30% aa identity, respectively, with the orthologous genes described in P. aeruginosa (Zhao, 1994). However, a higher identity of the PhhA gene product was observed with the phenylalanine hydroxylase of the betaproteobacterium Chromobacterium violaceum (Onishi, 1991) (Table 2). It should be noted that the PhhA protein of P. aeruginosa binds iron at the active site, like all mammalian aromatic amino acid hydroxylases (Zhao, 1994), whereas the C. violaceum PhhA protein is unique in its use of copper as a cofactor (Onishi, 1991), which suggests that the C. necator JMP134 PhhA gene product may also be a copper-containing enzyme. The following steps in the phenylalanine pathway in strain JMP134 are probably carried out by the products of the tyrB and hpd genes, which form a gene cluster together with aroP, that encode a general aromatic amino acid permease (C2 in Fig. 2. Table 2). This is different from most other proteobacteria, in which the hpd gene is not linked to other genes involved in phenylalanine and tyrosine degradation.

The homogentisate central pathway (Fig. 7) includes a homogentisate dioxygenase (HmgA) that opens the aromatic ring of homogentisate producing maleylacetoacetate which, in some bacteria, is directly hydrolyzed, yielding maleate and acetoacetate (Crawford, 1976); nevertheless, in most bacteria it is isomerized into fumarylacetoacetate (Chapman & Dagley, 1962). This isomerization is either catalyzed by a GSH-independent maleylacetoacetate isomerase, as in most Gram-positive bacteria (Hagedorn & Chapman, 1985), or by GSH-dependent enzymes (hmgC gene products), as has been reported in the Gram-negative strains D. acidovorans (Hareland, 1975), B. cepacia (Hamzah & Al-Baharna, 2001), and A. evansii KB740 (Mohamed Mel, 2002). Finally, fumarylacetoacetate is hydrolyzed by a specific hydrolase (HmgB) forming fumarate and acetoacetate (Fig. 7). Genes encoding enzymes of the homogentisate pathway are found scattered in the genome of strain JMP134 (Fig. 2, Table 2). A cluster comprising putative hmgA and hmgB1 genes is encoded in the small chromosome (Fig. 2), but a putative hmgC gene is located in a different region of the genome (C1 in Fig. 2). The nonlinkage of the hmgAB1 and hmgC genes has also been observed in R. solanacearum, Bordetella bronchiseptica, B. japonicum, Silicibacter pomeroyi and Pseudomonas syringae (Arias-Barrau, 2004). However, a second copy of a putative hmgB gene is found clustered to the hmgC gene (C1 in Fig. 2, Table 2). Both hmgB genes of C. necator JMP134 are equally related to the hmgB gene of P. putida U (Table 2), but the putative hmgB2 gene has only a 45% aa identity with hmgB1.

All three isomers of hydroxyphenylacetate are growth substrates for C. necator JMP134. Unfortunately, the information on the genes and biochemical functions involved in the catabolism of hydroxyphenylacetates in bacteria is too limited to search for the presence of such pathways in C. necator JMP134. However, it is possible to predict that, in this strain, at least 3- and 4-hydroxyphenylacetate would be catabolized by the homogentisate ring-cleavage pathway. 4-Hydroxyphenylacetate degradation has been reported to take place via homoprotocatechuate and a subsequent meta ring-cleavage pathway in the Gammaproteobacteria, E. coli and P. putida (Arunachalam, 1992; Olivera, 1994; Diaz, 2001). A genomic search for hpaBC genes that encode a two component 4-hydroxyphenylacetate-3-hydroxylase, and for an hpaD gene, that encodes a homoprotocatechuate-2,3-dioxygenase, does not show significant matches in the genome of strain JMP134; this indicates the absence of a meta ring-cleavage pathway for the degradation of homoprotocatechuate. A different 4-hydroxyphenylacetate degradation pathway has been reported in the Betaproteobacteria, D. acidovorans (Hareland, 1975), B. cepacia (Hamzah & Al-Baharna, 2001), and A. evansii KB740 (Mohamed Mel, 2002); it involves hydroxylation of the aromatic ring at C-1 with a concomitant migration of the carboxymethyl side chain to C-2 (the ‘NIH’ shift reaction), catalyzed by a NADH-dependent 4-hydroxyphenylacetate-1-hydroxylase and yielding homogentisate (Fig. 7). Therefore, it is likely that 4-hydroxyphenylacetate is metabolized by the homogentisate pathway in C. necator.

The degradation of 3-hydroxyphenylacetate has not been thoroughly studied. In E. coli, it has been reported that, like 4-hydroxyphenylacetate, this compound is degraded through the homoprotocatechuate pathway. However, in Flavobacterium species, a FAD-dependent 3-hydroxyphenylacetate-6-hydroxylase, that produces homogentisate, has been described and an N-terminal amino acid sequence has been determined (Van Berkel & Van Den Tweel, 1991). The use of this N-terminal sequence as an ‘in silico’ probe did not show any putative ORF in the C. necator JMP134 genome. It has been shown recently that P. putida U also metabolizes 3-hydroxyphenylacetate through the homogentisate pathway (Arias-Barrau, 2004), and genes mhaA and mhaB have been identified. They encode a 3-hydroxyphenylacetate-6-hydroxylase, a novel two-component flavoprotein aromatic hydroxylase. A mhaC gene that encodes a 3-hydroxyphenylacetate permease has also been identified (Arias-Barrau, 2004). In C. necator JMP134, a mhaA gene has been found that shows a 43% aa identity with the mhaA gene product from P. putida U (C2 in Fig. 2; Fig. 5, Table 2), but a mhaB homologous gene is absent.

The catabolism of 2-hydroxyphenylacetate has also been scarcely studied in bacteria. In early reports, 2-hydroxyphenylacetate was proposed as an intermediate in the phenylacetyl-CoA pathway, but recent studies discard this possibility in E. coli (Ismail, 2003). On the other hand, in P. fluorescens ST, it has been proposed that 2-hydroxyphenylacetate could be metabolized via homogentisate, based on the detection of a 2-hydroxyphenylacetate-5-hydroxylase activity that transforms 2-hydroxyphenylacetate into homogentisate in bacteria (Baggi, 1983), as has been reported for fungi (Mingot, 1999). This transformation is analogous to that catalyzed by salicylate-5-hydroxylase to produce gentisate in Ralstonia sp. U2 (Zhou, 2002). In strain JMP134, there are two putative gene clusters that would encode this enzyme (see ‘Degradation of salicylate and 3-hydroxybenzoate: the gentisate pathway’). One of these putative gene clusters has a much lower identity with the Ralstonia sp. U2 genes. Therefore, it is possible to speculate that this lower identity gene cluster encodes a 2-hydroxyphenylacetate-5-hydroxylase responsible for 2-hydroxyphenylacetate catabolism in C. necator JMP134.

The 2,3-dihydroxyphenylpropionate meta ring-cleavage pathway

The aerobic degradation of 3-hydroxyphenylpropionate (3-HPP) and 3-hydroxycinnamate is commonly started by a monooxygenase whose activity generates 2,3-dihydroxyphenylpropionate or 2,3-dihydroxycinnamate as central intermediates; these are further degraded via a meta ring-cleavage hydrolytic pathway (Diaz, 2001), that has been well described in E. coli (Ferrandez, 1997; Diaz, 1998, 2001; Torres, 2003). The 3-HPP and 3-hydroxycinnamate degradation pathway in E. coli is encoded by the mhp genes cluster. The MhpA monooxygenase transforms 3-HPP or 3-hydroxycinnamate into 2,3-dihydroxyphenylpropionate or 2,3-dihydroxycinnamate, respectively, which are then further converted into succinate (or fumarate, in the case of 3-hydroxycinnamate degradation), pyruvate, and acetyl-CoA, through the action of an extradiol dioxygenase (MhpB), a hydrolase (MhpC), a hydratase (MhpD), an aldolase (MhpE) and an acetaldehyde dehydrogenase (MhpF) (Fig. 3). A similar pathway has been described in C. testosteroni TA441, whose mhp gene cluster resembles that of E. coli (Arai, 1999b). In addition, a hpp gene cluster responsible for the partial catabolism of 3-HPP has been described in R. globerulus PWD1 (Barnes, 1997), but with a different gene organization, and a low sequence similarity with the mhp gene clusters of Gram-negative bacteria (Diaz, 2001). Cupriavidus necator JMP134 is able to grow on phenylpropionate, cinnamate, 3-HPP and 3-hydroxycinnamate, and a genomic search rendered a gene cluster resembling that of E. coli, but lacking the mhpE and mhpF genes, and with a slightly different gene organization (C1 in Fig. 2). The mhpE and mhpF genes encode the functions of the last two steps in the pathway: 4-hydroxy-2-ketovalerate aldolase and acetaldehyde dehydrogenase, respectively; these are shared with the catechol meta ring-cleavage pathway, which are encoded by the phlG and phlF genes in C. necator (Fig. 3). It is noteworthy that, in C. necator JMP134, phlG and phlF genes, thus named because of their higher identity to the gene products involved in the catechol meta ring-cleavage pathway (Table 3), are located apart from both gene clusters that encode meta ring-cleavage pathways (Fig. 2), unlike in gene organizations in other bacteria. In C. testosteroni TA441, in which both pathways have been described, there are two pairs of phlG and phlF genes located in the corresponding gene clusters (Arai, 1999b, 2000).

The C. necator JMP134 mhp genes have a moderate (50–70%) amino acid identity with the E. coli and C. testosteroni mhp genes (Table 3). The mhpA gene of strain JMP134 clusters together with other mhpA genes of Gram-negative bacteria and the hppA of R. globerulus PWD1, in the dendrogram of FAD-dependent hydroxylases (Fig. 5). It should be noted that a second putative FAD-dependent hydroxylase gene product (ReutB5808) of C. necator JMP134 groups with MhpA proteins (Fig. 5). However, it is unknown whether this gene product acts as a second 3-hydroxyphenylpropionate hydroxylase.

Cupriavidus necator JMP134 does not grow on 2-hydroxyphenylpropionate (melilotate) or 2-hydroxycinnamate, which could be explained by the inability of the MhpA protein to hydroxylate 2-hydroxyphenylpropionate to produce 2,3-dihydroxyphenylpropionate, as reported on the MhpA protein of E. coli (Burlingame & Chapman, 1983) and on the HppA protein of R. globerulus PWD1 (Barnes, 1997). In contrast, the strain Rhodococcus sp. V49 – which is able to grow on 2-hydroxyphenylpropionate using a 2,3-dihydroxyphenylpropionate meta ring-cleavage pathway – possesses a 2-hydroxyphenylpropionate hydroxylase encoded by the ohpB gene, that also belongs to the flavin-type aromatic hydroxylases family (Powell & Archer, 1998); nevertheless, its gene is more closely related to the monocomponent phenol hydroxylase encoding pheA gene of Pseudomonas sp. EST1001 (Nurk, 1991) than to the mhpA genes. No close relative of ohpB genes was found in the genome of strain JMP134, in concordance with the inability of this strain to grow on 2-hydroxyphenylpropionate or 2-hydroxycinnamate.

In C. necator JMP134, the 2,3-dihydroxyphenylpropionate-1,2-dioxygenase encoded by the mhpB gene, clusters with the type II extradiol dioxygenases (Fig. 9b) (Vaillancourt, 2004). The mhpB gene corresponds to the mpcI gene formerly described in C. necator JMP222 as a catechol-2,3-dioxygenase (Kabisch & Fortnagel, 1990a) (see ‘The catechol meta ring-cleavage pathway’). Later studies have shown that MpcI and MhpB proteins from E. coli are structurally and functionally related, and they have been proposed to constitute the type II family of extradiol dioxygenases (Spence, 1996). MpcI (MhpB) of C. necator JMP134 showed a broad specificity toward three-substituted catechols; propionate was found to be the optimum side chain (Spence, 1996). 2,3-Dihydroxycinnamate was as good a substrate as 2,3-dihydrophenylpropionate; this indicates that the enzyme is able to bind the alkyl side chain in a transoid conformation. Two other putative gene products in strain JMP134 (ReutB5775 and ReutB4784) have been assigned to type II extradiol dioxygenases, but they are distantly related to the MhpB gene product (Fig. 9b), and cluster with the extradiol dioxygenases involved in the degradation of syringate in Sphingomonas paucimobilis SYK-6 (Peng, 1998; Kasai, 2004).

The next step in this pathway is the hydrolytic cleavage of the extradiol ring fission product of 2,3-dihydrophenylpropionate; the resulting products are succinate (or fumarate from 2,3-dihydroxycinnamate) and 2-hydroxy-penta-2,4-dienoate (Fig. 3). This step is catalyzed by the 2-hydroxy-6-keto-nona-2,4-diene-1,9-dienoate hydrolase encoded by the mhpC gene. All other reported hydrolases that act on the ring-cleavage product of 2,3-dihydrophenylpropionate and 2,3-dihydroxycinnamate – i.e. MhpC from E. coli, MhpC from C. testosteroni TA441, HppC from R. globerulus PWD1, and OhpC from Rhodococcus sp. V49 – appear to be highly specific for the cleavage products (Barnes, 1997; Lam & Bugg, 1997; Powell & Archer, 1998; Arai, 1999a), so the same substrate specificity is expected for the MhpC gene product of C. necator. The hydrolase most closely related to MhpC of C. necator JMP134 is MhpC from E. coli. In contrast, the MhpD gene product from C. necator, responsible for the conversion of 2-hydroxy-penta-2,4-dienoate into 4-hydroxy-2-ketopentanoate, is most closely related to the MhpD protein of C. testosteroni TA441 (Table 3). The MhpD gene product is also homologous to the 2-hydroxypent-2,4-dienoate hydratase of the catechol meta ring-cleavage pathway in strain JMP134 (PhlE, 40% aa identity). A putative 3-hydroxyphenylpropionate transporter is encoded by the mhpT gene as part of the mhp genes cluster in C. necator; it shows a high amino acid identity with the MhpT protein of E. coli (Diaz, 2001), a member of the aromatic: H+ symporter family of transport proteins. Finally, a mhpR gene, divergently located in the C. necator mhp genes cluster (C1 in Fig. 2, Table 3), putatively encodes an IclR-type transcriptional regulator, which is homologous to the mhpR gene of E. coli (Torres, 2003). Although IclR-type regulators are generally transcriptional repressors, those which control catabolic pathways have always been described as activators (Tropel & van der Meer, 2004), including the MhpR protein of E. coli, which, in the presence of 3-HPP, activates the expression of mhp genes by binding to an operator region located upstream of the promoter (Torres, 2003).

Other catabolic pathways for aromatic compounds

The aerobic benzoyl-CoA pathway

A novel aerobic pathway for benzoate degradation has recently been described in A. evansii KB740. In this pathway (Fig. 10), benzoate is first converted, by an AMP forming benzoate-CoA ligase, into benzoyl-CoA. Benzoyl-CoA is hydroxylated and reduced at positions 2 and 3. The reaction is catalyzed by benzoyl-CoA oxygenase/reductase, a two component benzoyl-CoA dioxygenase, which is very dissimilar to other known oxygenase systems (Zaar, 2004), and it is composed by two proteins: an iron–sulfur-flavoprotein reductase (BoxA) and an oxygenase (BoxB). The dihydrodiol is the substrate for ring fission catalyzed by dihydrodiol lyase (BoxC) (Gescher, 2005). This homodimeric enzyme does not require oxygen and catalyzes the transformation to 3,4-dehydroadipyl-CoA semialdehyde. The latter intermediate is subsequently oxidized by 3,4-dehydroadipyl-CoA semialdehyde dehydrogenase (BoxD) to 3,4-dehydroadipyl-CoA (Gescher, 2006). The further metabolism is thought to lead to β-ketoadipyl-CoA, which is finally cleaved into succinyl-CoA and acetyl-CoA (Zaar, 2001). A cluster of fifteen genes that putatively encode this new benzoate pathway has been recently identified in the chromosome of A. evansii (Gescher, 2002). A gene cluster that putatively encodes a similar aerobic CoA-dependent pathway for benzoate degradation is also found in C. necator JMP134 (C1 in Fig. 2). This gene cluster comprises almost all the genes that are apparently essential for the benzoate catabolism in A. evansii (Table 4), including the benzoate CoA ligase (BzdA), the NADPH- and oxygen-dependent benzoyl-CoA oxygenase/reductase (BoxAB), the dihydrodiol lyase – involved in hydrolytic ring-cleavage (BoxC) – and a lactonase (ORF2), that putatively hydrolyzes the 3,6-lactone of the β-hydroxyadipyl-CoA formed in the last steps of the pathway (Fig. 10). Nevertheless, an 3,4-dehydroadipyl-CoA semialdehyde dehydrogenase (BoxD), which is involved in the oxidation of the aldehyde group in the 3,4-dehydroadipyl-CoA semialdehyde (Gescher, 2006), is absent. This enzyme activity can, however, be recruited from the dehydrogenase gene pool in C. necator JMP134. The C. necator's box gene cluster, like A. evansii's, includes an ABC transporter, putatively responsible for an effective benzoate uptake. However, whereas in A. evansii (Gescher, 2002) this ABC transporter comprises an ATP-binding membrane-spanning protein encoded by a single ORF (ORF4), C. necator contains two separate ORFs: one of these encodes the N-terminal domain (ORF4n) and the other the C-terminal domain (ORF4c) (not shown). A putative regulator, the BzdR gene product, with a high identity with the product of the bzdR gene of A. evansii CIB (Lopez Barragan, 2004) and with ORF10 of A. evansii KB740 (Gescher, 2002) is also encoded by the C. necator box gene cluster (Table 4). This regulator is a two-domain protein with an N-terminal domain that is similar to the regulatory proteins of the HTH family, and a C-terminus related to shikimate kinase I of E. coli. This suggests that the activity of this protein could be modulated by ATP-dependent phosphorylation in response to benzoate (Gescher, 2002).

The role of the box gene cluster in C. necator JMP134 is intriguing, because it is well established that strain JMP134 and related C. necator strains degrade benzoate through the β-ketoadipate pathway (Johnson & Stanier, 1971; Sauret-Ignazi, 1996; Ampe, 1997). The presence of two homologous copies of a gene cluster that encode for the enzymes of benzoate degradation via CoA activation, related to the box gene cluster of A. evansii, has been recently demonstrated in B. xenovorans LB400 (Gescher, 2002; Denef, 2004). Upregulation of one copy of these two gene clusters was found in cells grown on biphenyl, a compound that is degraded through benzoate, whereas no induction of these gene clusters was observed in benzoate grown cells (Denef, 2004). However, B. xenovorans LB400 mutants, which are defective in benzoate degradation via the β-ketoadipate pathway, were still able to grow on benzoate, recruiting both gene clusters for benzoate degradation via benzoyl-CoA (Denef, 2006). A similar situation may take place in C. necator JMP134, if the box gene cluster – putatively used for the degradation of the benzoyl-CoA produced from peripheral pathways – can be expressed when a high turnover of benzoate is required (because the β-ketoadipate pathway is defective) or if reduced oxygen tension is present, as has been suggested for B. xenovorans LB400 (Denef, 2006).

Pathways for amino- and nitroaromatic compounds

The 2-aminobenzoyl-CoA pathway

A new pathway for the metabolism of 2-aminobenzoate has been recently described in A. evansii KB740 (Schuhle, 2001). This not yet completely elucidated pathway (Fig. 10), similar to the benzoate pathway encoded by the box genes, begins with a 2-aminobenzoate-CoA ligase activity forming 2-aminobenzoyl-CoA; then, a 2-aminobenzoyl-CoA monooxygenase/reductase (ACMR) forms 2-amino-5-oxo-cyclohex-1-ene-1-carbonyl-CoA, and proceeds with β-oxidation (Schuhle, 2001). Two similar copies of the abm gene cluster – including the genes encoding 2-aminobenzoate-CoA ligase, the ACMR enzyme and three enzymes of a β-oxidation pathway – have been reported in A. evansii KB740 (Schuhle, 2001), and are coordinately expressed during the aerobic growth of this bacterium on 2-aminobenzoate. In C. necator, a gene cluster is found that encodes almost all the homologues of the A. evansii abm genes (C1 in Fig. 2, Table 4), with the exception of the abmF and abmH genes, which putatively encode a MarR-like regulator and a substrate-binding protein of an ABC transporter, respectively (Schuhle, 2001). In addition, a gene (abmX) that putatively encodes a thioesterase – with similarity to one found in the box gene cluster of A. evansii (see previous section) – is also present in the abm gene cluster of C. necator (Table 4). This putative thioesterase can undertake the final steps of the β-oxidation pathway in C. necator, involved in 2-aminobenzoate catabolism. It is worth noting that in the closely related betaproteobacterium B. cepacia DBO1 (Chang, 2003), and in the two Gammaproteobacteria Pseudomonas resinovorans (Urata, 2004) and A. baylyi ADP1 (Eby, 2001), a ‘classical anthranilate pathway’ has been described; in it, anthranilate is converted into catechol by anthranilate-1,2-dioxygenase and metabolized through the classical β-ketoadipate pathway. A genomic search in strain JMP134 does not yield any similar ORF. In addition, an early study in the closely related strain C. necator 335 showed no induction of the enzymes of the β-ketoadipate pathway in tryptophan-grown cells (Johnson & Stanier, 1971), which excludes the possibility that 2-aminobenzoate is metabolized by this pathway. In C. metallidurans CH34, a three-step pathway of aerobic l-tryptophan degradation to anthranilate has been described recently (Kurnasov, 2003). The kynBAU operon encoding three required enzymes for this pathway: tryptophan 2,3-dioxygenase (gene kynA), kynurenine formamidase (gene kynB), and kynureninase (gene kynU), is found in C. necator JMP134 (Table 4). Taken together, these clues indicate that the kyn and abm gene clusters in C. necator are responsible for tryptophan catabolism through a 2-aminobenzoyl-CoA pathway.

The 3-hydroxyanthranilate pathway

A pathway for the degradation of 2-nitrobenzoate has been recently described in P. fluorescens KU-7 (Hasegawa, 2000; Muraki, 2003). This pathway is peculiar, because it proceeds through the meta ring-cleavage of 3-hydroxyanthranilate, an aromatic intermediate which has been described before in the kynurenine pathway of eukaryotic organisms, but not in bacteria (Kucharczyk, 1998). Further catabolism of 3-hydroxyanthranilate is undertaken by a 3-hydroxyanthranilate-3,4-dioxygenase (NbaC), which cleaves the aromatic ring into 2-amino-3-carboxymuconate-6-semialdehyde, while the 2-amino-3-carboxymuconate-6-semialdehyde decarboxylase (NbaD) catalyzes the decarboxylation of the latter compound into 2-aminomuconate-6-semialdehyde (Fig. 3). The subsequent action of 2-aminomuconate-6-semialdehyde dehydrogenase (NbaE), 2-aminomuconate deaminase (NbaF), 4-oxalocrotonate decarboxylase (NbaG), 2-oxopent-4-dienoate hydratase (NbaH), 4-hydroxy-2-oxovalerate aldolase (NbaI) and acylating aldehyde dehydrogenase (NbaJ), finally produce pyruvate and acetyl-CoA. The nba genes responsible for the 3-hydroxyanthranilate meta ring-cleavage pathway in P. fluorescens KU-7 have been sequenced (Muraki, 2003), and a gene cluster closely resembling that of P. fluorescens KU-7 has been found in C. necator (C2 in Fig. 2). However, C. necator JMP134 is unable to grow on 2-nitrobenzoate and the corresponding genes for 3-hydroxyanthranilate meta ring-cleavage pathway have been designated as haa genes (3-hydroxyanthranilic acid). The haa gene cluster of C. necator shows amino acid sequence identities ranging from 64% to 72% (Table 3), excluding the LysR-type regulator haaR gene, which shows a lower identity with nbaR. The haa gene cluster encodes all the genes that are essential for the catabolism of 3-hydroxyanthranilate, with the exception of nbaI and nbaJ genes. However, these two functions are shared with the catechol- and the 2,3-dihydroxyphenylpropionate meta ring-cleavage pathways of C. necator (see ‘The catechol meta ring-cleavage pathway’ and ‘The 2,3-dihydroxyphenylpropionate meta ring-cleavage pathway’) and can be recruited from the above mentioned PhlG and PhlF gene products.

Catabolic pathways for nitrophenols

Three nitrophenols support the growth of C. necator JMP134: 2,6-dinitrophenol (Ecker, 1992), 3-nitrophenol (Schenzle, 1997) and 2-chloro-5-nitrophenol (Schenzle, 1999a). The initial degradation pathway for 3-nitrophenol consists of the transformation into 3-hydroxylaminophenol and then into aminohydroquinone; these are catalyzed by a 3-nitrophenol nitroreductase (MnpA) and a 3-hydroxylaminophenol mutase (Fig. 11), respectively (Schenzle, 1997, 1999b). The 3-hydroxylaminophenol mutase was identified as a glutamine syntethase (Table 5) (Schenzle, 1999b). The initial degradation of 2-chloro-5-nitrophenol is analogous to that of 3-nitrophenol, and results in the formation of 2-amino-5-chlorohydroquinone (Schenzle, 1999a). The removal of the chlorine group in the latter intermediate is reductively mediated and produces aminohydroquinone, a common intermediate. Further degradation of aminohydroquinone (G. Zylstra, pers. commun.) (Fig. 11), proceeds through the ring-cleavage between the adjacent hydroxyl and amino groups and is catalyzed by an aminohydroquinone dioxygenase (MnpC); then, the amide group is removed by an amidase (MnpD) to form maleylacetate, which is transformed into β-ketoadipate by a maleylacetate reductase (MAR) (MnpE). It is worth mentioning that, of the six MAR genes present in C. necator (Fig. 12), mnpE gene is the less related one (see next sections). A gene cluster, mnpCARED (megaplasmid pJPL in Fig. 2), that putatively encodes the enzymes that catalyze the conversion of 3-nitrophenol into β-ketoadipate is found in strain JMP134. The amino acid sequences of the Mnp proteins of C. necator have low identity levels with known homologues (Table 5). Interestingly, putative gene sequences are found at the flanks of the mnpCARED gene cluster, that encode the dehalogenating activity in 2-chloro-5-nitrophenol degradation (mnpF); and an aminobenzoquinone reductase activity that forms aminohydroquinone (mnpG).

Figure 11

The (amino)hydroquinone and (chloro)hydroxyquinol pathways.

View this table:
Table 5

Genes encoding pathways for (chloro)hydroxyquinol, (amino)hydroquinone and methylhydroquinone and peripheral reactions

GenePosition (bp)No. aaRelated gene products
NameFunction/descriptionOrganism% Id (aa)Accession no.References
mnpFpJPL 108024-106981347linD2,5-Dichlorohydroquinone reductive dechlorinaseSphingomonas paucimobilis UT2637 (342)P95806Miyauchi (1998)
mnpHpJPL 108969-108079296RSc0758Putative oxygenase oxidoreductase proteinRalstonia solanacearum GMI100033 (258)NP_518879Salanoubat (2002)
mnpDpJPL 110515-109109468amdAEnantiomer-selective amidaseRhodococcus sp.33 (455)AAA26183Mayaux (1991)
mnpEpJPL 111627-110509372tcbFMaleylacetate reductasePseudomonas sp. P5131 (273)AAD13629Muller (1996)
mnpRpJPL 112657-111746303RSp0662Probable transcription regulator, LysR familyRalstonia solanacearum GMI100040 (296)NP_522223Salanoubat (2002)
mnpApJPL 112842-113528228nbzANitrobenzene nitroreductasePseudomonas putida HS12 pNB171 (219)AAK26512Park & Kim (2000)
mnpCpJPL 113641-114591316linEHydroquinone meta-cleavage dioxygenaseSphingomonas paucimobilis UT2640 (316)AAQ96752Miyauchi (1998)
mnpGpJPL 114613-115239208orf1Quinone reductasePseudomonas pseudoalcaligenes JS4576 (199)AAT71309Chae and Zylstra (unpublished data)
glnA C1 2261396-2259981471glnAGlutamine synthetaseHerbaspirillum seropedicae Z7883 (471)AAC32389Persuhn (2000)
hqoD C2 1393905-1392847352tftEMaleylacetate reductaseBurkholderia cepacia AC110062 (351)Q45072Daubaras (1995)
hqoEC2 1395607-1394648319linEHydroquinone meta-cleavage dioxygenaseSphingomonas paucimobilis UT2643 (317)Q9WXE6Miyauchi (1998)
hxqD C2 768585-767512357pnpDMaleylacetate reductaseRalstonia sp. SJ9869 (352)AAS87585Pandey et al. (unpublished data)
hxqCC2 777451-776546301hadCHydroxyquinol-1,2-dioxygenaseRalstonia pickettii DTP060265 (263)BAA13107Hatta (1999)
tcpRC1 1707124-1706153323pcpRPCP degradation regulatory proteinSphingobium chlorophenolicum36 (294)P52679Cai & Xun (2002)
tcpX C1 1707275-1707835186tftCChlorophenol-4-monooxygenase component 1Burkholderia cepacia AC110053 (168)AAC23547Hubner (1998)
tcpA C1 1707954-1709507517hadAChlorophenol-4-hydroxylaseRalstonia pickettii DTP060286 (517)BAA13105Hatta (1999)
tcpB C1 1709570-1710160196hadBNADH dehydrogenase/NAD(P)H reductaseRalstonia pickettii DTP060276 (190)BAA13106Hatta (1999)
tcpC C1 1710178-1711008276hadCHydroxyquinol-1, 2-dioxygenaseRalstonia pickettii DTP060272 (274)BAA13107Hatta (1999)
tcpYC1 1711124 1712086320ebA5757Protein involved in regulation of phenol degradationAzoarcus sp. EbN134 (296)CAI09392Rabus (2005)
tcpD C1 1712101-1713168355macAMaleylacetate reductaseRalstonia eutropha 335T65 (359)AAD55886Seibert (2004)
mhqBC2 1372734-1373696320mhqBExtradiol dioxygenaseBurkholderia sp. NF10057 (301)BAE46530Tago (2005)
mhqCC2 1373785-1374639284dntGHydrolaseBurkholderia cepacia R3456 (281)AAL50014Johnson (2002)
mhqAC2 1374676-1376454592mhqAMonooxygenaseBurkholderia sp. NF10061 (595)BAE46529Tago (2005)
mhqRC2 1376539-1377321260bphRTranscriptional regulator, GntR-familyBurkholderia xenovorans LB40036 (212)P37335Erickson & Mondello (1992)
  • * Genes for those cases where genetic or biochemical evidence for function is available are underlined.

Figure 12

Dendrogram showing the relatedness of maleylacetate reductases. The dendrogram was obtained by the neighbor-joining method using mega 4.0 based on sequence alignments calculated by clustal w using the default options. Sequences of deduced proteins encoded in the genome of Cupriavidus necator JMP134 are highlighted black. The sequences and their accession numbers are as follows: HqoD JMP134, C. necator JMP134 maleylacetate reductase (YP_298887); TftE AC1100, Burkholderia cepacia AC1100 maleylacetate reductase (AAC43333); HxqD JMP134, C. necator JMP134 maleylacetate reductase (YP_298327); PnpD SJ98, Ralstonia sp. strain SJ98 maleylacetate reductase (AAS87585); TcpD JMP134, C. necator JMP134 maleylacetate reductase (YP_295799); MacA 335, C. necator 335 maleylacetate reductase (AAD55886); MacA 1CP, Ralstonia opacus 1CP maleylacetate reductase (AAC38802); DxnE RW1, Sphingomonas sp. strain RW1 maleylacetate reductase (ZP_01610140); GraC MTP-10005, Rhizobium sp. strain MTP-10005 maleylacetate reductase (BAF44524); ScaC SA1, Novosphingobium subarcticum SA1 maleylacetate reductase (AAW29743); ClpF S1, Defluvibacter lusatiensis S1 maleylacetate reductase (CAD60257); TfdF2 TFD44, Sphingomonas sp. strain TFD44 maleylacetate reductase (AAT99371); TfdF TFD44, Sphingomonas sp. strain TFD44 maleylacetate reductase (AAT99365); TfdF 2A, B. cepacia 2A maleylacetate reductase (AAK81685); TfdF EST4002, Achromobacter xylosooxidans ssp. denitrificans EST4002 maleylacetate reductase (AAS49435); TfdFI JMP134, C. necator JMP134 maleylacetate reductase (YP_025384); TfdF NK8, Burkholderia sp. strain NK8 maleylacetate reductase (BAB56012); ClcE B13, Pseudomonas knackmussii B13 maleylacetate reductase (AAB71540); TetF RW71, Pseudomonas chlororaphis RW71 maleylacetate reductase (CAB89825); TcbF P51, Pseudomonas sp. strain P51 maleylacetate reductase (AAD13629); TfdF P4a, Delftia acidovorans P4a chloromaleylacetate reductase (AAK57010); CphF-I A6, Arthrobacter chlorophenolicus A6 maleylacetate reductase (AAO46999); NpdC JS444, Arthrobacter sp.` JS444 maleylacetate reductase, G. Zylstra unpublished data; MnpE JMP134, C. necator strain JMP134 maleylacetate reductase (YP_293177); PnpE ENV2030, Pseudomonas sp. ENV2030 maleylacetate reductase, G. Zylstra unpublished data; PnpE JS443, Pseudomonas putida JS443 maleylacetate reductase, G. Zylstra unpublished data; TfdFII JMP134, C. necator JMP134 maleylacetate reductase (YP_025392); LinF UT26, Sphingobium japonicum UT26 maleylacetate reductase (BAD66863); PcpE ATCC 39723, Sphingobium chlorophenolicum ATCC 39723 chloromaleylacetate reductase (AAM96664); PcpE UG30, Sphingobium sp. strain UG30 2-chloromaleylacetate/maleylacetate reductase (AAS20423).

The (chloro)hydroxyquinol ring-cleavage pathway

With the exception of 2,4,6-trichlorophenol (2,4,6-TCP), and 2,4-dichlorophenol, C. necator is unable to use chlorinated phenols as carbon sources (Clement, 1995). Whereas 2,4-dichlorophenol is metabolized via 3,5-dichlorocatechol (see ‘Tfd functions: biochemistry and genetics’), 2,4,6-TCP, – like 2,4,5-trichlorophenol (2,4,5-TCP) in other bacteria – is metabolized through the so-called (chloro)hydroxyquinol pathway (Fig. 11) (Kasberg, 1995; Zaborina, 1995). The first step, catalyzed by the 2,4,6-TCP monooxygenase (TcpA), is the oxidative conversion of 2,4,6-TCP into 2,6-dichlorobenzoquinone, followed by a hydrolytic dechlorination that produces 6-chloro-2-hydroxybenzoquinone (Xun & Webster, 2004). 6-chloro-2-hydroxybenzoquinone is either chemically or enzymatically reduced (probably by TcpB) to 6-chloro-2-hydroxybenzoquinol (Fig. 11). Contrary to previous assumptions, 2,6-dichlorobenzoquinol is not an intermediate of 2,4,6-TCP degradation in strain JMP134 (Xun & Webster, 2004). 6-Chloro-2-hydroxybenzoquinol is transformed into 2-chloromaleylacetate by 1,2-hydroxyquinol dioxygenase (TcpC), and then converted into β-ketoadipate by MAR (TcpD) (Louie, 2002; Matus, 2003). This pathway is different from that reported for 2,4,5-TCP degradation in B. cepacia AC1100, which starts with the oxidation of 2,4,5-TCP to 2,5-dichlorobenzoquinone, and its further transformation into 5-chloro-2-hydroxybenzoquinol (Kasberg, 1995). The latter compound is dechlorinated to hydroxybenzoquinone and then reduced to hydroxybenzoquinol by a quinone reductase. Thus, all chloride substituents are removed from the aromatic ring before its cleavage. Whereas the hydroxyquinol-1,2-dioxygenase of strain AC1100 is unable to use 5-chloro- or 6-chlorohydroxyquinol (Kasberg, 1995), hydroxyquinol-1,2-dioxygenases of 2,4,6-TCP-degrading strains usually use chlorohydroxyquinol and hydroxyquinol as substrates, although they vary in their substrate preferences (Kasberg, 1995; Latus, 1995; Zaborina, 1995; Hatta, 1999). (Chloro)hydroxyquinol dioxygenases form a distinct group in the dendrogram of intradiol-1,2-dioxygenases (Fig. 4), that also includes the enzyme from Arthrobacter sp. strain BA-5-17 (Murakami, 1999).

In strain JMP134, enzymes for the metabolism of 2,4,6-TCP are encoded by the tcpRXABCYD gene cluster (C1 in Fig. 2) (Matus, 2003). Sequence analysis indicates that tcpA and tcpB genes encode a FADH2 utilizing monooxygenase – with a 65% aa sequence identity with the TftD protein from B. cepacia AC1100 – and a flavin reductase, respectively (Table 5). However, the analysis of mutants of strain JMP134 that are defective in different tcp genes, showed that tcpB is not required for the conversion of 2,4,6-TCP (Louie, 2002; Sanchez & Gonzalez, 2007). It has been shown very recently that TcpB has activity for quinone reduction with FMN or FAD as the cofactor, and NADH as the reductant (Belchik & Xun, 2008). Sequence comparison with the tftC gene – which encodes a flavin reductase in B. cepacia AC1100 – strongly suggests that, as in 2,4,5-TCP degradation, a NADH: FAD oxidoreductase is involved in the initial monooxygenation and that in strain JMP134 this function is carried out by the TcpX gene product (Fig. 11, Table 5) (Matus, 2003; Sanchez & Gonzalez, 2007). In vitro assays of coupling of TcpX and TcpA demonstrated that TcpX provided FADH2 for TcpA catalysis (Belchik & Xun, 2008).

Fluorobenzoate catabolism

4-Fluorobenzoate degradation is common to most Cupriavidus strains (Schlomann, 1990b). The degradation of 4-fluorobenzoate does not require the enzymes specialized in halocatechol degradation (see next section), as evidenced by the fact that C. necator JMP222 – a derivative of strain JMP134 cured of plasmid pJP4 – grows on 4-fluorobenzoate (Schlomann, 1990b). The degradation pathway for this halobenzoate, studied in the strain C. necator 335, is started by the transformation of 4-fluorobenzoate into 4-fluorocatechol, performed by benzoate dioxygenase and benzoate dihydrodiol dehydrogenase [see ‘The ortho ring-cleavage pathways for benzoate and p-hydroxybenzoate (the β-ketoadipate central pathway)]. In contrast to chlorinated derivatives (see next section), 4-fluorocatechol is a good substrate for proteobacterial catechol-1,2-dioxygenases [see ‘The ortho ring-cleavage pathways for benzoate and p-hydroxybenzoate (the β-ketoadipate central pathway)] which produce 3-fluoromuconate. Cycloisomerization of this ring-cleavage product results in the formation of 4-fluoromuconolactone (Schlomann, 1990a) (see next section). In both Cupriavidus strains, JMP134 and 335, a trans-dienelactone hydrolase that produces maleylacetate has been reported to be induced during growth in 4-fluorobenzoate (Schlomann, 1990b); this enzyme is supposed to transform 4-fluoromuconolactone into maleylacetate (Nikodem, 2003). The pathway is completed with the transformation of maleylacetate into β-ketoadipate by MAR. Recent evidence indicates that in strain 335, a MAR encoding gene, macA, is found in a gene cluster along with macB gene, which hypothetically encodes a membrane transport protein, and macR gene – which encodes a putative regulator (Seibert, 2004). The fact that strain JMP222 is able to grow on 4-fluorobenzoate is indicative that plasmid-encoded MAR would be functional for maleylacetate turnover, but not essential. Any of the chromosome-encoded MAR would assume the main role in 4-fluorobenzoate degradation, being tcpD, hqxD and hqoD genes more closely related to macA gene than mnpE gene (Fig. 12), and probably are functionally redundant. It should be noted that no ORF similar to the macR gene from strain 335 is found in the neighborhood of the tcpD, hqxD or hqoD genes, or elsewhere in the genome of strain JMP134.

Catabolic pathway for mono- and dichlorinated compounds: the tfd genes

Tfd functions: biochemistry and genetics

The catabolic pathway for 2,4-D has been thoroughly studied in strain JMP134 (Fig. 13). This pathway is encoded by tfd (two, four-dichlorophenoxyacetate) genes (Fig. 13b), which are located in the pJP4 plasmid, and initiated by a 2,4-D/α-ketoglutarate dioxygenase (Fukumori & Hausinger, 1993a, b). The same pathway has been reported in various 2,4-D-degrading isolates (Beadle & Smith, 1982; Chaudhry & Huang, 1988; Bhat, 1994; Maltseva, 1996; Poh, 2002; Thiel, 2005); however, in some strains, 2,4-D degradation is started by monooxygenases, labeled CadAB, that are related to the TftAB protein involved in the degradation of 2,4,5-trichlorophenoxyacetic acid (Kitagawa, 2002). Both catabolic activities form 2,4-dichlorophenol as intermediate. In the pathway initiated by the TfdA protein, α-ketoglutarate is transformed into succinate and CO2, and 2,4-D is converted into glyoxylate and 2,4-dichlorophenol. The enzyme of strain JMP134 uses several other phenoxyacetates as substrates: MCPA, phenoxyacetate, 2-methylphenoxyacetate, 4-methylphenoxyacetate and 2-chlorophenoxyacetate (Pieper, 1988); however, not all of these compounds support the growth of the wild type strain (Pieper, 1988, 1989). The inability to use these phenoxyacetates as growth substrates is not due to a restricted specificity of the TfdA protein but to regulatory constraints, as mutants that constitutively express tfd genes could grow on all these compounds (Fig. 3) (Pieper, 1989). Even 2-naphthoxyacetate, benzofuran-2-carboxylate or 2,4-dichlorocinnamate are substrates for the TfdA protein of strain JMP134 (Dunning Hotopp & Hausinger, 2001). Based on the ability of the TfdA protein to metabolize chlorinated cinnamic acids, it has been proposed that tfdA-like sequences present in 2,4-D-nondegrading bacteria may metabolize substituted cinnamic acids (Dunning Hotopp & Hausinger, 2001).

Figure 13

The tfd genes encoded pathways (a) and organization (b).

The next step in 2,4-D degradation is the transformation of 2,4-dichlorophenol into 3,5-dichlorocatechol (Fig. 13a), which is catalyzed by the 2,4-dichlorophenol hydroxylase, a single component flavoprotein monooxygenase (TfdB). In pJP4, there are two genes that encode chlorophenol hydroxylases (Fig. 13b), named tfdBI and tfdBII. The enzyme activity previously purified from C. necator JMP134 grown on 2,4-D, corresponds to TfdBI (Liu & Chapman, 1984; Farhana & New, 1997), but the substrate profile of TfdBII is similar (Ledger, 2006): both enzymes use 2,4-dichlorophenol, 4-methyl-2-chlorophenol and 2- and 4-monosubtituted phenols as substrates, and exhibit only poor activity with phenol (Liu & Chapman, 1984; Farhana & New, 1997; Ledger, 2006). Moreover, both gene sequences map in the same branch of the dendrogram for FAD-dependent hydroxylases (Fig. 5).

TfdA and TfdB proteins transform 2,4-D, via 2,4-dichlorophenol, into 3,5-dichlorocatechol (3,5-DCC), a central intermediate in chloroaromatic metabolism. The reactions that produce the corresponding chlorocatechols during growth on 3-CB are carried out by (1) the benzoate dioxygenase, which introduces two oxygen atoms into the benzoate molecule to produce the benzoate-cis-1,2-dihydrodiol derivative, and (2) the benzoate dihydrodiol dehydrogenase, which restores aromaticity by forming the catechol (Fig. 13a) (Pieper, 1993). These enzymes are recruited from the benzoate degradation pathway and exhibit some activity with 3-chlorobenzoate, but not with 4-chlorobenzoate [see ‘The ortho ring-cleavage pathways for benzoate and p-hydroxybenzoate (the β-ketoadipate central pathway)’]. It is interesting to note that these enzymes produce 70% of 3-chlorocatechol (3-CC) and 30% of 4-chlorocatechol (4-CC) from 3-CB (Pieper, 1993).

The chlorocatechols produced during growth on 2,4-D or 3-CB are metabolized through the chlorocatechol ortho ring-cleavage pathway (Fig. 13a). This pathway has been reported in a number of chloroaromatic-degrading bacteria (Reineke, 1998; Timmis & Pieper, 1999; Pieper & Reineke, 2000). In C. necator JMP134, like in other bacteria, the chlorocatechol ortho ring-cleavage pathway is initiated by chlorocatechol-1,2-dioxygenase (TfdC), which produces 2,4-dichloromuconate, 3-chloromuconate and 2-chloromuconate from 3,5-DCC, 4-CC, and 3-CC, respectively (Fig. 13a). Two genes that encode TfdC (Fig. 13b) are present in strain JMP134, and both TfdCI and TfdCII proteins show a similar substrate profile (Perez-Pantoja, 2000; Plumeier, 2002). However, under the same growth conditions, the activity of TfdCI enzyme is about two to three times higher than that of TfdCII enzyme. Whether this is due to higher enzyme amounts or to differences in their specific activity remains to be elucidated. It should also be mentioned that chlorocatechol-1,2-dioxygenases usually exhibit a broad substrate range, whereas catechol-1,2-dioxygenase, only transforms 4-chlorocatechol at significant rates (Pieper, 1993).

Chloromuconates formed by intradiol ring-cleavage of chlorocatechol are substrates for chloromuconate cycloisomerase (TfdD), which converts them into the corresponding dienelactones (Vollmer & Schlomann, 1995). Dechlorination was shown to be enzyme catalyzed, as muconate cycloisomerases form 2- and 5-chloromuconolactone from 2-chloromuconate and are not able to dechlorinate (Vollmer, 1994). In contrast, chloromuconate cycloisomerases form trans-dienelactone through a specific cycloisomerization to 5-chloromuconolactone and the rotation of the catalytic center of the lactone ring that allows proton abstraction and thus dehalogenation (Schell, 1999). In the case of 3-chloromuconate cycloisomerization by muconate cycloisomerases, a highly unstable intermediate, 4-chloromuconolactone, is first formed; then dechlorination produces protoanemonin. Only chloromuconate cycloisomerases form cis-dienelactone, probably via an enol-enolate intermediate (Pieper & Reineke, 2004).

As in the case of tfdC genes, there are also two copies of tfdD genes (Fig. 13b). The TfdDI enzyme produces dienelactones from 2-chloro-, 3-chloro- and 2,4-dichloromuconate at significant rates; this substrate profile is shared by other chloromuconate cycloisomerases from Gram-negative strains (Vollmer, 1999). In contrast, the TfdDII enzyme exhibits high activity against 3-chloromuconate (forming cis-dienelactone) and poor activity against 2-chloromuconate. Moreover, this enzyme's activity results in an equilibrium between 2-chloromuconate and 5-chloro- and 2-chloromuconolactone, and it is very inefficient in catalyzing dehalogenation to form trans-dienelactone; therefore, it differs from all (chloro)muconate cycloisomerases described so far (Plumeier, 2002).

Dienelactones are converted into maleylacetates by dienelactone hydrolase, and MAR catalyzes the reduction of the double bond of the maleylacetate to form β-ketoadipate, the common metabolite of the catechol and the chlorocatechol ortho ring-cleavage pathways (Kaschabek & Reineke, 1992). There are two tfdE genes (Fig. 13b); however, the activity levels of the TfdEI protein, with cis-dienelactone as a substrate, are significantly higher than those of the TfdEII protein (Plumeier, 2002). Maleylacetates with chlorine substituents in the 2-position, such as 2-chloromaleylacetate (formed from 3,5-dichlorocatechol as intermediate of 2,4-D metabolism), are reduced by MAR in a first step to yield maleylacetate. Obviously, the enzymatic attack on the C2-carbon results in an intermediate that spontaneously eliminates chloride. The tfd gene clusters encode two tfdF genes (Fig. 13b), which means that the genome of C. necator has six MAR encoding genes (see previous sections and Fig. 12). The presence of such level of gene redundancy is intriguing; it is of key importance to study the expression of these genes in C. necator cells exposed to compounds whose degradation pathway requires MAR activity. Further degradation of the β-ketoadipate formed by MAR proceeds as in the catechol ring-cleavage pathway, using pcaIJF gene encoded functions (see ‘The pob and pca genes’); however, it is also possible that the related functions encoded in the mml genes (see ‘The methylcatechol ortho ring-cleavage pathway’) also play a role in chlorocatechol degradation.

The overall organization of tfd genes, which are located in a 22-kb region of the pJP4 plasmid and have two copies of the chlorocatechol ortho ring-cleavage pathway genes (Fig. 13), is unique among the 2,4-D-degrading strains, although D. acidovorans P4a has also been reported to comprise two tfd gene clusters (Hoffmann, 2003). The tfd-I gene cluster of strain JMP134 encodes one putative LysR-type regulator (the TfdT gene product), which is interrupted in its carboxyl end by an ISJP4 insertion sequence (Leveau & van der Meer, 1997). The tfd-II cluster encodes the same functions as the tfd-I cluster; however, the tfdDII and tfdCII genes are in a different order. Moreover, this cluster comprises the tfdK gene, which encodes a membrane protein involved in 2,4-D transport (Leveau, 1998) (Fig. 13a). The tfdA gene is found close to this gene cluster together with two inverted, perfect copies of the LysR-type regulatory genes: tfdR and tfdS (Fig. 13b). The gene organization of the tfd-I gene cluster is similar to that observed for the clc genes of P. putida AC25 (Ghosal & You, 1988, 1989) and Pseudomonas knackmussii B13 (Frantz, 1987), the tcb genes of Pseudomonas sp. P51 (van der Meer, 1991), the cbn genes of C. necator NH9 (Ogawa & Miyashita, 1999), and the tfd genes of Burkholderia sp. NK8 (Liu, 2001) and D. acidovorans P4a (Hoffmann, 2003). All these gene clusters – except for that of Burkholderia sp. NK8 and of the strain JMP134 – are characterized by the presence of an ORF of unknown function between the genes that encode chloromuconate cycloisomerase and dienelactone hydrolase. In strain JMP134, the tfd-II gene cluster organization is similar to the tfdRCEBKA gene order of the 2,4-D-degrading strains Variovorax paradoxus TV1 (AB028643), B. cepacia 2a (Poh, 2002), D. acidovorans P4a (Hoffmann, 2003) and Achromobacter xylosooxidans ssp. denitrificans EST4002 (Vedler, 2004). However, the tfd-II cluster of strain JMP134 is the only one that comprises the tfdD and tfdF genes. No clear trend in the evolutionary relatedness of the tfd counterparts is observed in strain JMP134. Whereas the tfdB genes map relatively close in the corresponding dendrogram (Fig. 5), the tfdC genes (Fig. 4), and specially the tfdF genes, are clearly part of unrelated branches (Fig. 12). This suggests that these two clusters are not the product of a recent gene duplication event. It is possible that the presence of redundant gene clusters in strain JMP134 may be the effect of a lateral acquisition of catabolic genes (Trefault, 2004).

What would be the role of these two gene clusters in strain JMP134? Although each tfd gene cluster is enough to allow growth on 3-CB (Perez-Pantoja, 2000; Plumeier, 2002), some reports suggest that the catabolic functions encoded by the tfd-II gene cluster are not required (Don, 1985; Laemmli, 2004). However, the presence of two apparently redundant gene clusters would be important for the ‘fine tuning’ of the expression of catabolic functions. On the one hand, the number of copies of these tfd gene clusters is important for the catabolic performance of these strains (Klemba, 2000; Trefault, 2002), and for the adequate response to the toxicity of catabolic intermediates, e.g. chlorocatechols (Perez-Pantoja, 2003). On the other hand, the presence of two tfdB encoded functions would be important to prevent the accumulation of the toxic intermediate 2,4-dichlorophenol (Ledger, 2006).

Regulation of the tfd genes

One of the most interesting aspects of the tfd catabolic genes, their regulation, is still not well understood. Its study has been affected by the recent realization that tfd gene organization is complex and most gene functions are redundant. Available evidence indicates that all the tfd genes are expressed during growth on 2,4-D (Leveau, 1999; Laemmli, 2004). However, it should be emphasized that studies with 3-CB or other substrates that are metabolized through the chlorocatechol ring-cleavage pathway are required, because it is highly possible that the tfd genes are expressed in a growth-substrate depending manner. Early reports indicate that two regulatory genes control the expression of the tfd genes: tfdS, described as an activator of the expression of the tfdA gene (You & Ghosal, 1995) and a repressor of the tfdB gene (Kaphammer & Olsen, 1990), and tfdR, reported to regulate the tfd-I gene cluster (Harker, 1989). After the work of Matrubutham & Harker (1994), which showed that the tfdS and tfdR genes were identical, earlier reports needed to be revised. The use of constructs with different copy numbers and different hosts have been proposed to explain these early results (Leveau & van der Meer, 1996). The role of the tfdR gene as a regulatory element of the tfd-I gene cluster has been nicely shown by J.R. van der Meer's lab (Leveau & van der Meer, 1996), which has provided evidence that the tfdR gene can replace the ISJP4-interrupted tfdT gene. The role of tfdR gene as a master regulatory gene for tfd gene expression is one of the few aspects that are clear. Three intergenic regions –tfdT-tfdCI, (tfdT/C), tfdR-tfdDII (tfdR/D) and tfdA-tfdS (tfdA/S) (Fig. 13b) – share significant levels of nucleotide sequence identity (40–60%) with the intergenic regions involved in (chloro)catechol catabolism that are regulated by other LysR-type transcriptional activators (Matrubutham & Harker, 1994; McFall, 1998). Evidence has been found for in vitro binding of the TfdR protein to the tfdT/C region (Matrubutham & Harker, 1994), which coincides with its role as activator of the tfd-I genes cluster. Albeit indirect, additional, support for the regulatory role of the TfdR protein on both tfd gene clusters is provided by the fact that the tfd-I and tfd-II modules, cloned separately and under the control of the tfdR gene, express all the corresponding Tfd enzymes (Perez-Pantoja, 2000). Despite this evidence, the role of the TfdR protein in the expression of the tfdA gene is still unclear. Preliminary evidence indicates, however, that the TfdR protein is able to bind to the tfdA/S intergenic region (N. Trefault, L. Guzmán, M. Manzano, D.H. Pieper & B. González, unpublished data), but further investigation is clearly required.

Is there any differential role for the TfdS and the TfdR proteins? Although the idea that two identical genes would produce proteins with different activities is bizarre, the effect of their respective positions along with very limiting amounts of a regulatory protein may produce a differential effect on gene expression of the immediately adjacent intergenic regions (tfdA/S for the TfdS protein and tfdR/D for the TfdR protein). tfdS and tfdR-inactivated pJP4 derivatives may help to explore this point. In this respect, it should be noted that a pJP4 derivative lacking the tfdS-tfdR region (pYG1010) is able to support growth on 3-CB, but not on 2,4-D (You & Ghosal, 1995). This could lead to the conclusion that tfdS/tfdR gene regulation is not necessary for the expression of tfd genes during growth on 3-CB (You & Ghosal, 1995), but a cross-talk effect mediated by catR chromosomal functions, such as those present in C. necator [see ‘The ortho ring-cleavage pathways for benzoate and p-hydroxybenzoate (the β-ketoadipate central pathway)’] has also been suggested as an explanation (Leveau & van der Meer, 1996). Although plausible, such cross-talk needs to be demonstrated. In addition, a possible participation of the truncated TfdT protein cannot be fully discarded, because the binding of the TfdT protein to the tfdT/C intergenic region has been demonstrated (Leveau & van der Meer, 1996). It is important to note that the tfdT gene of strain JMP134 has a significant level of identity with the tfdT gene of the tfd gene cluster of Burkholderia sp. NK8 (Liu, 2001). This TfdT protein responds to chloromuconates, chlorocatechols and even chlorobenzoates, an inducer profile which is very different from other LysR-type regulators involved in (chloro)catechol catabolism. This makes possible that the TfdT protein from strain JMP134 may play an unexpected role in regulation of tfd gene expression.

Which are the inducers involved in the LysR-type gene mediated regulation of the tfd genes? By analogy with the catechol pathway, the corresponding (chloro)muconates may be the inducers for the chlorocatechol ortho ring-cleavage pathway. This has been clearly shown by Chakrabarty's lab for clc genes and cbn genes (McFall, 1997c; Ogawa, 1999). Their studies have proven that ClcR can bind two sequences in the clcA upstream region, and that this binding is modified by the presence of 2-chloromuconate and muconate. This shift in binding of the ClcR protein to clcA DNA is necessary for transcriptional activation. Such studies have also shown that the CatR protein interacts in a slightly different way than the CIcR protein with the catA promoter region (it can bind to three sequences, in a dimer/tetramer fashion). The CatR protein only interacts with muconate (McFall, 1997b) and the CatR-mediated transcriptional activation is not repressed by fumarate (McFall, 1997a), as has been reported for the ClcR protein.

Little is known about the tfd genes system. A genetic approach was used in order to identify the inducer of tfd gene expression (Filer & Harker, 1997). Because only mutants that were blocked in the tfdDI and tfdEI genes showed an increased level of induction in the presence of 2,4-D – whereas mutants blocked in tfdA, tfdBI or tfdCI genes did not – 2,4-dichloromuconate was suggested as the inducer. Considering the presence of a second tfd gene module, these observations are difficult to explain, inasmuch as only mutants blocked in the tfd-I genes were used in this work. In fact, the TfdCII protein (Plumeier, 2002) should allow further metabolism of 3,5-dichlorocatechol in the tfdCI mutant. However, as the relative contribution of the first three enzymes from the tfd-I gene cluster is two to five times higher than those from the tfd-II gene cluster (Perez-Pantoja, 2000), the overall balance may favor the functions encoded by tfd-I gene cluster. This explanation may also be useful to understand why transposon mutagenesis of tfd-I genes produced an accumulation of catabolic intermediates (Don, 1985). In this context, the possibility that the TfdR protein interacts with 2,4-dichloromuconate to control the tfdA gene expression is interesting, but has not yet been proved.

The genetic background of C. necator JMP134 is clearly important for an adequate tfd gene expression. Interestingly, 26 out of 28 different betaproteobacterial strains harboring pJP4 grow efficiently on 2,4-D, whereas 17 out of 20 different alpha- or gammaproteobacterial strains harboring pJP4 do not (D. Pérez-Pantoja, B. González, unpublished data). This observation suggests that tfd gene expression clearly requires chromosomally encoded functions that are usually present in Betaproteobacteria, but absent in alpha- or gammaproteobacterial strains.

The pJP4 plasmid

The broad-host, conjugative, IncPβ plasmid pJP4, has recently been sequenced (Trefault, 2004). The tfd genes are flanked by two IS1071 elements in the well-conserved IncPβ backbone. IS1071 elements have been associated with several gene clusters that encode the degradation of anthropogenic compounds (Sota, 2006). ISJP4 elements (Leveau & van der Meer, 1997) and a complex transposon, Tn5504 (Trefault, 2004), are also present in this catabolic plasmid. There are about 20 other ORFs in the non IncPβ backbone region of the plasmid, but none of them is directly involved in aromatic compound degradation (Trefault, 2004).

Several genetic changes in the pJP4 plasmid have been reported; for example, 40 kb deletions, including the tfd genes region, have been observed after transposon mutagenesis with Tn1771 (Don, 1985). Tn10-stimulated pJP4 deletions in a noncatabolic region have been also described (Clement, 2000). Early work described large genetic changes in the pJP4 plasmid, after transfer of the pJP4 plasmid to P. putida and a selection for growth on 3-CB (Ghosal, 1985). Electron microscopy, Southern analyzes and the restriction enzyme profile of the obtained pJP4 plasmid derivative (pYG2) clearly suggested that a c. 15 kb deletion, along with c. 25 kb duplication had taken place. About 15 years later, another pJP4 plasmid derivative, pJP4-F3 – obtained by subculturing C. necator JMP134 in liquid cultures containing 3-CB – appeared to undergo the same rearrangement as the pYG2 plasmid. The molecular characterization of pJP4-F3 plasmid showed (Clement, 2001) that the 15-kb deletion is flanked by the tfdS/tfdR region and the IS1071 insertion sequence. This implies a loss of the tfdA gene and explains the inability to grow on 2,4-D that is observed in strains harboring pYG2 or pJP4-F3 plasmids. The duplication produced a 21-kb inverted repeat that included the tfdS/R region and all the tfd genes, except tfdA. A plausible model for this pJP4 plasmid rearrangement has been proposed (Clement, 2001): it consists of an intermolecular, double crossover, homologous recombination. Some pJP4 plasmid features predicted by this model, as the presence of two sequences related to the IS1071, were later confirmed by the complete sequencing of the plasmid (Trefault, 2004). The main features of this rearrangement on pJP4 plasmid, at the molecular and plasmid population level, have been recently studied using a multiple-PCR approach (Larrain-Linton, 2006). It is clear that, in wild-type populations, the pJP4 plasmid form is the most abundant, but that at least two recombinant forms – pJP4-F3 and pJP-FM – are also detectable (around 1% of plasmid population). Successive transfers of the wild type in 3-CB strongly select cells harboring the pJP4-F3 plasmid form, which is the derivative with a higher tfd gene dosage (Clement, 2001), but this enrichment can be reversed after transfers in 2,4-D. In this context, it is worth mentioning that a C. necator JMP134 derivative with the integrated, one copy per cell, pJP4 plasmid, does not grow on 3-CB (Trefault, 2002). Given that the pJP4 plasmid forms appear in about five copies per cell (Trefault, 2002), the presence of only one copy can be detrimental for growth on chloroaromatic compounds. Two is a threshold number of copies that allows the clc element to support growth on chlorobenzene (Ravatn, 1998), which suggests that small differences in gene dosage may have a critical effect on growth properties. As indicated in ‘Regulation of the tfd genes’, the avoidance of chlorocatechol and chlorophenol toxicity requires an adequate balance between the producing and the transforming activities of chlorocatechol and chlorophenol (Perez-Pantoja, 2003; Ledger, 2006).

A model for the pJP4 plasmid evolution has recently been proposed (Trefault, 2004); it suggests the simultaneous acquisition of both tfd gene clusters in the IncPβ backbone of the pJP4 precursor (Trefault, 2004). Several lines of evidence support this possibility. Among them, the fact that only simultaneous acquisition of both tfd gene clusters allows the bacterium to grow efficiently on 3-CB or 2,4-D, avoiding toxicity problems. The unique tfd gene organization in pJP4 may have arisen from the complementation between the two tfd gene modules and the concerted regulation commanded by the activity of the TfdR protein.

Concluding remarks

Cupriavidus necator possesses 11 of the 12 main routes for aromatic degradation reported in Proteobacteria; the only one absent is the homoprotocatechuate pathway. Functional redundancy seems to be a key feature in the ability of strain JMP134 to degrade a significant number of different aromatic compounds. Redundant functions were observed in the catechol, protocatechuate, salicylate and phenylacetyl-CoA pathways; in the degradation of benzoate and chloroaromatic compounds; in some of the 4-hydroxybenzoate and (methyl)phenols peripheral reactions; and in the presence of several meta ring-cleavage enzymes and other oxygenases, maleylacetate reductases and regulatory proteins. Interestingly, the genome of C. necator encodes more than 70 oxygenases (Table 6) that belong to the main oxygenase groups reported for aromatic compound catabolism. The systematic metabolic reconstruction work reviewed here could only assign functions to half of these oxygenases. Whether the unknown oxygenase functions are involved in aromatic compounds degradation remains to be studied.

View this table:
Table 6

Oxygenases related to catabolism of aromatic compounds encoded in the genome of Cupriavidus necator JMP134

Oxygenase familyNo.Gene products in C. necator JMP134
Intradiol dioxygenases8CatA1, CatA2, TfdCI, TfdCII, TcpC, HxqC, PcaGH, ReutB5855
Type I extradiol dioxygenases7PhlB, MhqB, ReutC6234, ReutA1133, ReutB5807, ReutB5647, ReutA1557
Type II extradiol dioxygenases3MhpB, ReutB4784, ReutB5775
Type III extradiol dioxygenases4MhbD1, MhbD2, HmgA, HaaC
Hydroquinone dioxygenases2MnpC, HqoE
Tryptophan-2,3-dioxygenases2MnpH, KynA
Rieske-type [2Fe–2S] dioxygenases, α subunit13BenA, NagG, ReutB3776, ReutA1473, ReutB5781, ReutB4795, ReutC6324, ReutB5789, ReutB4340, ReutD6483, ReutB4815, ReutA0879, ReutB5836
Multicomponent di-iron monooxygenases, α subunit5PhlN1, PhlN2, TbcA, PaaA1, PaaA2
Single-component flavoprotein monooxygenases15PobA, PobB, MhbA1, MhbA2, TfdBI, TfdBII, MhqA, MhpA, MhaA, ReutB3601, ReutA2515, ReutB5808, ReutC6230, ReutB4218, ReutB5646
Two-component flavoprotein monooxygenases5TcpA, ReutB5779, ReutA1542, ReutC6314, ReutB3469
α-Ketoglutarate-dependent dioxygenases3TfdA, ReutB5293, ReutC6361
4-Hydroxyphenylpyruvate dioxygenases3Hpd, ReutB5028, ReutB5035
Pterin-dependent monooxygenases1PhhA
Cytochrome P450 monooxygenases2ReutB5278, ReutB5115
Benzoyl-CoA oxygenases1BoxA
2-Aminobenzoyl-CoA monooxygenases2AbmA, ReutB5482

Is the high catabolic versatility of strain JMP134 common to other soil bacteria? Genome-wide studies performed on P. putida KT2440 (Nelson, 2002); B. xenovorans LB400 (Chain, 2006), Rhodococcus sp. strain RHA1 (McLeod, 2006), and ‘A. aromaticum’ sp. EbN1 (Rabus, 2005), show a similar level of catabolic versatility; this clearly suggests that this kind of bacteria may be more common in nature than previously expected. Are most of these catabolic gene clusters evolutionary remnants, or an adaptative improvement in the constant competition for limited carbon sources in natural habitats? A plausible assumption would be that bacteria, such as strain JMP134, were naturally exposed to mixtures of different aromatic compounds (in low amounts), that could be used as carbon sources; for example, during microbial degradation of lignin, exposure to plant exudates or in environments polluted with aromatic compound mixtures. If such was the case, would all the catabolic genes be coordinately expressed? In other words, are aromatic compounds hierarchically degraded? Are there catabolic misroutings when mixtures of aromatic compounds, metabolized by different pathways, are being used as carbon sources? Studies of the gene expression profile using catabolic DNA microarrays will clearly help to answer such questions. Obviously, classical biochemical work on key enzymes would be needed to fully understand the catabolism of aromatic compound mixtures. The analysis of protein expression profiles would also help to compare the relative contribution of different gene clusters during growth on mixtures of aromatic compounds.


This work has been funded by the FONDECYT grants 1030493, and 1070343, the ‘Millennium Institute for Fundamental and Applied Biology’, the ‘Millennium Nucleus in Microbial Ecology and Environmental Microbiology and Biotechnology’ and the ICA4-CT-2002-10011 (ACCESS) Contract of the European Union. Joint Genome Institute work was performed under the auspices of the United States Department of Energy's Office of Science, Biological and Environmental Research Program and by the University of California, Lawrence Livermore National Laboratory under Contract No. W-7405-Eng-48, Lawrence Berkeley National Laboratory under contract No. DE-AC02-05CH11231 and Los Alamos National Laboratory under contract No. DE-AC02-06NA25396. D.P.-P. is a CONICYT-DAAD Ph.D. fellow. R.D.I. is a MECESUP Ph.D. fellow.


  • Editor: Alexander Boronin