Genetics and Molecular Biology

Genetic Analysis of Phenoxyalkanoic Acid Degradation in Sphingomonas herbicidovorans MH

Tina A. Müller, Steven M. Byrde, Christoph Werlen, Jan Roelof van der Meer, Hans-Peter E. Kohler
DOI: 10.1128/AEM.70.10.6066-6075.2004

ABSTRACT

Phenoxyalkanoic acid degradation is well studied in Beta- and Gammaproteobacteria, but the genetic background has not been elucidated so far in Alphaproteobacteria. We report the isolation of several genes involved in dichlor- and mecoprop degradation from the alphaproteobacterium Sphingomonas herbicidovorans MH and propose that the degradation proceeds analogously to that previously reported for 2,4-dichlorophenoxyacetic acid (2,4-D). Two genes for α-ketoglutarate-dependent dioxygenases, sdpAMH and rdpAMH, were found, both of which were adjacent to sequences with potential insertion elements. Furthermore, a gene for a dichlorophenol hydroxylase (tfdB), a putative regulatory gene (cadR), two genes for dichlorocatechol 1,2-dioxygenases (dccAI/II), two for dienelactone hydrolases (dccDI/II), part of a gene for maleylacetate reductase (dccE), and one gene for a potential phenoxyalkanoic acid permease were isolated. In contrast to other 2,4-D degraders, the sdp, rdp, and dcc genes were scattered over the genome and their expression was not tightly regulated. No coherent pattern was derived on the possible origin of the sdp, rdp, and dcc pathway genes. rdpAMH was 99% identical to rdpAMC1, an (R)-dichlorprop/α-ketoglutarate dioxygenase from Delftia acidovorans MC1, which is evidence for a recent gene exchange between Alpha- and Betaproteobacteria. Conversely, DccAI and DccAII did not group within the known chlorocatechol 1,2-dioxygenases, but formed a separate branch in clustering analysis. This suggests a different reservoir and reduced transfer for the genes of the modified ortho-cleavage pathway in Alphaproteobacteria compared with the ones in Beta- and Gammaproteobacteria.

Phenoxyalkanoic acid herbicides such as 2,4-dichlorophenoxyacetic acid (2,4-D), chiral mecoprop [(R,S)-2-(4-chloro-2-methyl-phenoxypropanoic acid)], and chiral dichlorprop [(R,S)-2-(2,4-dichlorophenoxypropanoic acid)] (Fig. 1) are widely used against broadleaved weeds in agriculture, lawn pastures, and industries. They were introduced in large amounts into the environment in the 1940s and 1950s (1, 10, 57). 2,4-D, mecoprop, and dichlorprop are synthetic compounds and have only been applied for some decades. However, microorganisms able to use them as the sole carbon and energy source have been isolated from different environments (21, 36, 52, 53, 60). 2,4-D-degrading strains from the Beta- and Gammaproteobacteria groups mostly harbor similar genes for 2,4-D degradation (17, 24, 34, 50, 51), which supports that frequent gene exchange has occurred and hence their adaptation and wide distribution (17).

FIG. 1.
FIG. 1.

Pathway proposed for dichlorprop degradation in S. herbicidovorans MH and for 2,4-D degradation in R. eutropha JMP134(pJP4) (30, 39-41). The enzymes of the pathway in S. herbicidovorans MH are printed in boldface. The enzymes of the genes, which have not yet or partially been isolated from S. herbicidovorans MH are displayed within parentheses.

In many cases, the first step in the degradation of phenoxyalkanoic acids is the enzymatic conversion of the acids to the corresponding phenols, then via the catechols, and through the modified ortho-cleavage pathway (21, 43). The best-studied degradation pathway is that of 2,4-D in the soil bacterium Ralstonia eutropha JMP134(pJP4) (Fig. 1). Both the genetic and the enzymatic details of the pathway in this bacterium have been established. R. eutropha JMP134(pJP4) degrades 2,4-D by an α-ketoglutarate-dependent dioxygenase that cleaves the ether bond to produce 2,4-dichlorophenol (15, 16, 45). 2,4-Dichlorophenol in turn is hydroxylated to 3,5-dichlorocatechol by a phenol-hydroxylase (TfdBI/II) (13, 29, 40). 3,5-Dichlorocatechol undergoes ortho-ring fission to 2,4-dichloro-cis,cis-muconate, a reaction catalyzed by chlorocatechol 1,2-dioxygenase (TfdCI/II) (29, 39-41). 2,4-Dichloro-cis,cis-muconate is then converted to cis-2-chlorodienelactone and further to 2-chloromaleylacetate by chloromuconate cycloisomerase (TfdDI/II) and dienelactone hydrolase activity (TfdEI/II), respectively (27, 40). Finally, chloromaleylacetate is reduced to 3-oxoadipate via maleylacetate by maleylacetate reductase (TfdFI/II) (40, 41). Tett et al. (48) showed that (R)-mecoprop degradation in Alcaligenes denitrificans proceeds analogously to 2,4-D degradation in R. eutropha JMP134(pJP4). Similarly, phenol hydroxylase, chlorocatechol 1,2-dioxygenase, and chloromuconate cycloisomerase activities are induced in Delftia acidovorans MC1 upon exposure to dichlorprop. This suggests that dichlorprop is also metabolized through a modified ortho-cleavage pathway (6).

Sphingomonas herbicidovorans MH is a versatile phenoxyalkanoic acid degrader and grows on mecoprop and dichlorprop as the sole carbon and energy sources (22, 26, 60). Although strain MH metabolizes both enantiomers, it preferentially uses the (S) enantiomer. Nickel et al. (38) showed that S. herbicidovorans MH harbors two distinct α-ketoglutarate-dependent dioxygenase activities that are involved in the enantioselective degradation of mecoprop and dichlorprop to achiral phenols. However, nothing is known about the genetic background of the mecoprop and dichlorprop degradation pathway in strain MH. S. herbicidovorans MH belongs to the Alphaproteobacteria, whereas other well-studied phenoxyalkanoic acid degraders such as R. eutropha JMP134(pJP4) and D. acidovorans MC1 belong to the Betaproteobacteria and the Gammaproteobacteria. Total DNA from phenoxyalkanoic acid-degrading Alphaproteobacteria did not hybridize to tfdA gene probes encoding α-ketoglutarate-dependent 2,4-D dioxygenase (17, 24, 34, 50, 51). PCR experiments aimed at amplifying DNA fragments from Sphingomonas strains with tfdA primers also failed (44, 50). Therefore, the hypothesis was formulated that Alphaproteobacteria might harbor different dioxygenases catalyzing the initial step of phenoxyalkanoic acid degradation. Only recently, the first tfdA homologous gene, tfdAα, was isolated from a 2,4-D-degrading bacterium belonging to the Bradyrhizobium-Agromyces-Nitrobacter-Afipia (BANA) cluster of Alphaproteobacteria (23). On the other hand, fragments similar to phenol hydroxylase (e.g., tfdB like) and chlorocatechol 1,2-dioxygenase genes (e.g., tfdC like) have also been amplified from Sphingomonas strains with degenerate primers in the PCR (31, 50). Whereas the tfdB-like gene fragments from Sphingomonas strains were more than 60% similar to tfdB genes from Beta- and Gammaproteobacteria, tfdC-like gene fragments showed little similarity to other known chlorocatechol 1,2-dioxygenase genes and formed a rather coherent group among themselves (50). Hence, Alphaproteobacteria might not only harbor different dioxygenases but also different chlorocatechol degradation genes, and it was thus proposed that gene flow might be less common between Alpha- and Beta- or Gammaproteobacteria than among Beta- and Gammaproteobacteria themselves (17, 18).

In order to test this hypothesis and to elucidate the phenoxyalkanoic acid degradation pathway, we isolated and characterized several genes from S. herbicidovorans MH by PCR amplification with degenerated primers, DNA-DNA hybridization, cosmid library construction, and DNA sequencing. We propose that these genes are involved in phenoxyalkanoic acid degradation in strain MH and are equivalent to those known from the 2,4-D degradation pathway of R. eutropha JMP134(pJP4). The expression of the characterized genes was analyzed in hybridization experiments with mRNA isolated from S. herbicidovorans MH cultures that were induced with mecoprop or 2,4-D. The two chlorocatechol 1,2-dioxygenases were expressed in Escherichia coli and characterized in terms of their substrate specificities. Finally, the phylogenetic relationships of the S. herbicidovorans MH chlorocatechol pathway genes were compared to those of other phenoxyalkanoic acid degradation pathways.

MATERIALS AND METHODS

Bacterial strains and culture conditions.S. herbicidovorans MH was grown in baffled Erlenmeyer flasks shaking at 30°C either in complex medium (59) or in mineral medium supplemented with the appropriate carbon and energy source. The mineral medium was prepared as described in Nickel et al. (38) and was modified by reducing the amount of added peptone to 10 mg/liter and by adding 10-mg/liter yeast extract. E. coli strains were grown at either 30 or 37°C in Luria-Bertani (LB) medium (42) with the appropriate antibiotic. Ampicillin and kanamycin were added to final concentrations of 50 μg/ml, and chloramphenicol was added to 25 μg/ml. If necessary, 5-bromo-4-chloro-3-indolyl-β-d-galactoside (Biosynth AG, Staad, Switzerland) was added to a final concentration of 50 μg/ml.

Standard molecular techniques.Cloning and digestions were done according to established procedures (4, 42). Restriction enzymes and other DNA-modifying enzymes were purchased from Promega (Wallisellen, Switzerland) and Fermentas (Nunningen, Switzerland). Plasmids and cosmids were isolated by the boiling Miniprep method, the alkaline lysis method according to Sambrook et al. (42), or with the E.Z.N.A. plasmid Miniprep kit II (Peqlab Biotechnologies GmbH, Baden-Dättwil, Switzerland) as suggested by the manufacturer. Gel extraction was carried out with the MinElute gel extraction kit (QIAGEN AG, Basel, Switzerland) according to the protocol of the supplier.

Construction of a cosmid library of S. herbicidovorans MH.The SuperCos1 cosmid vector kit (Strategene, Amsterdam, The Netherlands) was used to construct a cosmid library. S. herbicidovorans MH was grown in 500 ml of mineral medium containing 200-mg/liter (R,S)-mecoprop until a turbidity of 0.15 (measured at 546 nm) was reached. The culture was harvested by centrifugation for 30 min at 6,000 × g. The pellet was resuspended in 15 ml of lysis buffer (10 mM NaCl, 20 mM Tris-HCl, pH 8, 1 mM EDTA, 100-μg/ml proteinase K, and 0.5% [wt/vol] sodium dodecyl sulfate) and incubated overnight at 50°C. The DNA was isolated by phenol-chloroform extraction (42), precipitated with ethanol, and dissolved in 4 ml of TE (10 mM Tris-HCl, pH 8, 1 mM EDTA) and stored at 4°C. Partial digestion of the DNA was done with Sau3AI using different enzyme concentrations for 15 min at 37°C. The digestion was stopped by adding EDTA to a final concentration of 1 mM. The preparation of the host strain E. coli XL-1 BlueMR, the packaging, and the titer determination were carried out according to the protocol of the supplier (Stratagene). Finally, about a thousand clones were picked and grown overnight in microtiter plates in LB medium supplemented with kanamycin at 30°C. Glycerol was added to a final concentration of 15%, and the cosmid library was stored at −80°C.

Screening for genes involved in phenoxyalkanoic acid degradation by PCR.To isolate rdpA, we amplified a fragment with the PCR primers RDPin_fG and RDPin_r (55) from chromosomal DNA of S. herbicidovorans MH. Oligonucleotides for PCR were obtained from Microsynth GmbH (Balgach, Switzerland). The following conditions were applied: initial denaturation for 5 min at 93.5°C; denaturing for 30 s at 93.5°C; annealing for 30 s at 52, 49, 46, 43, 41, and 39°C; extension for 45 s at 72°C (5 cycles at each annealing temperature, 10 cycles at 39°C) followed by a final extension for 4 min at 72°C. A chlorophenol hydroxylase gene fragment (tfdB) was amplified with the tfdBup and tfdBlow primers from total DNA of S. herbicidovorans MH as described by Vallaeys et al. (50).

To screen for a chlorocatechol 1,2-dioxygenase gene, the degenerate PCR primers CCDb and CCDe described by Leander et al. (31) were used. The PCR protocol was as follows: initial denaturation for 6 min at 93.5°C, 35 cycles of denaturation for 1 min at 95°C, annealing for 1 min at 48°C, and extension for 1 min at 72°C followed by one extension cycle for 6 min at 72°C. PCR fragments were cloned in pGEM-T Easy (Promega) and sequenced to check the insert (Table 1).

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TABLE 1.

Plasmids constructed in this work

Hybridization of the cosmid library and construction of plasmids harboring genes involved in phenoxyalkanoic acid degradation.For colony blotting of the cosmid library, the colonies were grown overnight on LB agar plates at 37°C. Colony material was transferred to Hybond-XL membrane (Amersham Biosciences, Dübendorf, Switzerland) by direct contact. The colony material on the membrane was lysed twice with a solution of 0.5 M NaOH for 2 min and washed twice with a solution of 1 M Tris-HCl (pH 8). The DNA was fixed on the membrane by UV light (254 nm, 120 mJ/cm2) in a Stratalinker model 1800 (Stratagene).

The cosmid library of S. herbicidovorans MH was hybridized with the inserts of pMec1 (dccA fragment) and pMec5 (rdpA fragment). The fragments were radioactively labeled with [α-32P]dATP using the Random Primed DNA labeling kit according to the supplier's protocol (Roche Applied Science, Rotkreuz, Switzerland). Positively reacting cosmids were isolated, digested, and checked by Southern hybridization. Additionally, all positive-reacting cosmids were hybridized in the same way with the insert of pMec23 to screen for the tfdB gene. Suitable fragments were then subcloned in pUC vectors (5), yielding the plasmids listed in Table 1.

Transposon insertion and sequencing.The EZ::TN<KAN-2> Tnp transposome kit (Epicentre Technologies, Madison, Wis.) was used for the insertion of a kanamycin resistance marker and for sequencing priming sites into the plasmids with cloned S. herbicidovorans MH DNA. The insertion reaction was carried out according to the protocol of the supplier. DNA sequencing was performed on double-stranded DNA templates with the Thermo Sequenase primer cycle sequencing kit with 7-deaza-dGTP (Amersham Biosciences) and M13 primers (5) or KAN-2 FP-1/RP-1 primers (Epicentre Technologies). Sequencing reactions were analyzed on an automated DNA sequencer model 4200 IR2 (LI-COR, Inc., Lincoln, Nebr.). Primers for sequencing were labeled at the 5′ end with IRD-700 or IRD-800 and were purchased from MWG Biotech AG (Ebersberg, Germany). Sequencing data were analyzed with DNASTAR software (DNASTAR, Inc., Madison, Wis.) and compared to the databases by BLAST searches (www.ncbi.nlm.nih.gov/BLAST) (47).

Construction of expression plasmids pMec9 and pMec21, expression of DccAI/II in E. coli, and preparation of cell extract.The dccAI gene was reamplified from a cosmid fragment by PCR with the primer pair CCD2/CCD3 (5′-GAGAGGTCATATGAGCAATCGT-3′, 5′-AACGCGAGGATCCGCAGACCAT-3′), thereby introducing an NdeI restriction site and a BamHI restriction site (underlined). The dccAII gene was amplified by PCR with the forward primer ClcA2_for (5′-AGGTTCCATGGGCAATCG-3′) to introduce an NcoI site and the reverse primer ClcA2_rev (5′-AGGATCCGTGCCTGTCAG-3′) to create a BamHI site. The fragments were first cloned into pGEM-T Easy and verified by sequencing. They were then digested with the respective enzymes and cloned in the same restriction sites in pRSET6α and pET8c, respectively (46) (Table 1).

To express the proteins, a preculture of E. coli BL21(DE3)(pLysS) harboring pMec9 or pMec21 was grown overnight at 37°C. A volume of 1% (vol/vol) of the preculture was transferred to fresh medium, and the cells were grown at 30°C until a turbidity of 0.4 to 0.6 (measured at 546 nm) was reached, after which isopropyl-β-d-thiogalactopyranoside was added to a final concentration of 0.05 mM. The culture was incubated for another 3 h at 30°C, and the cells were harvested by centrifugation. A pellet of approximately 1 g (wet weight) was resuspended in 2.5 ml of a solution of 40 mM Tris-HCl, pH 7.4, containing 0.3 mM EDTA. The cells were broken by one passage through a French Pressure cell at 1 MPa. The lysate was centrifuged at 16,000 × g for 30 min at 4°C, and the supernatant was recovered and stored at −20°C until used as cell extract in enzyme activity measurements.

Enzyme assay.Enzymatic activity of chlorocatechol 1,2-dioxygenase with all substrates was measured spectrophotometrically at 260 nm except for protocatechuate, which was monitored at 290 nm. To 485 μl of buffer (containing 40 mM Tris-HCl, 0.3 mM EDTA, pH 7.4), 10 μl of cell extract of E. coli BL21(DE3)(pLysS) harboring pMec9 or pMec21 containing between 15 and 30 μg of total protein was added. The reaction was started by adding the substrate from a 20 mM stock in methanol to a final concentration of 0.4 mM. To calculate activity, extinction coefficients were taken from Broderick and O'Halloran (7) and Dorn and Knackmuss (11).

Induction experiment.S. herbicidovorans MH was grown on mineral medium with 20 mM pyruvate to a turbidity of 0.4 to 0.6 (measured at 546 nm). The culture was induced by the addition of (R)-, (S)-mecoprop, or 2,4-D dissolved in dimethyl sulfoxide (DMSO; 10 g/liter) to a final concentration of 100 mg/liter and incubated for another 30 min at 30°C. A culture to which only DSMO was added was used as a negative control. To stop induction and prevent RNA degradation, 1 volume of culture was added to 2 volumes of RNA protector immediately after sampling (QIAGEN AG). The mixture was incubated at room temperature for 5 min, centrifuged at 13,000 × g for 1 min, and the supernatant was decanted. The pellets were stored at −20°C.

RNA extraction.RNA was extracted with the RNeasy Mini kit from QIAGEN AG according to the supplier's protocol. The RNA, dissolved in RNase-free water, was then stored by adding 2 volumes of ethanol and 0.1 volume of 5 M sodium acetate (pH 5.2) at −20°C. RNA concentrations were determined spectrophotometrically at 260 nm.

In vitro synthesis of antisense mRNA.The pGEM-T Easy plasmids pMec14, -18, and -20, harboring rdpA, sdpA, and dccAII, respectively, were linearized by restriction enzyme digestion in order to allow insert-specific transcription from either the T7 or the SP6 promoter. In vitro transcription reactions were carried out with biotin-16-UTP and T7 or SP6 RNA polymerase with the Riboprobe in vitro transcription systems (Promega) according to the protocol of the supplier. The size and yield of all antisense mRNA probes were checked on 1% agarose gels.

Dot blot hybridization of mRNA.Total RNAs were washed with 70% ethanol and dissolved in 50 μl of RNase-free water. Equal volumes of RNA dilutions (0.5, 0.1, and 0.05 μg per 50 μl) were blotted onto positively charged nylon membranes (QIAGEN GmbH) in a dot blot manifold (Invitrogen Life Technologies, Basel, Switzerland) containing a 96-well 3-mm gasket. The samples were washed with an equal volume of 1× SSC (0.15 M NaCl, 15 mM sodium citrate) directly after their application. On each blot, a series of DNA standards were included (not shown). The standards contained 100, 50, 10, 1, or 0.1 ng of DNA/50 μl of plasmid pMec14, -18, or -20.

Phylogenetic analysis.Amino acid sequences were aligned with the software tool CLUSTALX (49). Phylogenetic analysis was done with the maximum-likelihood method by using the software Tree-Puzzle (http://www.tree-puzzle.de).

Chemicals.(R,S)-, (R)-mecoprop, and 2,4-D were purchased at Riedel-de Haën (Buchs, Switzerland; PESTANAL, 99% purity). (S)-Mecoprop had been synthesized previously by Zipper (58). Catechol, 4-chloro-, 3,5-dichloro-, 4,5-dichloro-, tetrachloro-, 3-methyl-, and 4-nitrocatechol were purchased from Sigma-Aldrich (Buchs, Switzerland; 97 to 99% purity). 3-Chlorocatechol was bought at Promochem GmbH (Wesel, Germany; 99% purity). All other chemicals were either obtained from Fluka (Buchs, Switzerland) or from Merck AG (Dietikon, Switzerland).

Nucleotide sequence accession number.Nucleotide sequences determined in this study were deposited in the EMBL database under accession no. AJ628859 to AJ628863.

RESULTS

Sequence determination of sdpA, rdpA, and tfdB.Degradation of phenoxyalkanoic acid herbicides usually begins with an ether cleavage to the corresponding phenol by α-ketoglutarate-dependent dioxygenases (15, 16, 45, 55, 56). By using a primer set previously developed for isolating a gene fragment encoding a (R)-dichlorprop dioxygenase from D. acidovorans MC1 (55), a 0.3-kb fragment was amplified from total DNA of S. herbicidovorans MH. This PCR-amplified fragment was cloned (pMec5) and sequenced to confirm its identity to the dichlorprop dioxygenase gene. It was subsequently used to screen a cosmid library of S. herbicidovorans MH. Several cosmids were positively hybridizing with the pMec5 insert, and a 2.8-kb NcoI-fragment was identified from cosmid 10A12 to contain the region of identity to pMec5. This fragment was subcloned (pMec10), sequenced, and analyzed in detail (Table 2). The sequence revealed two open reading frames (ORFs), one of which was almost identical to the (R)-dichlorprop dioxygenase gene rdpA from D. acidovorans MC1 (rdpAMC1; Table 2 and Fig. 2A). At the nucleotide level, the 888-bp rdpA ORF from S. herbicidovorans MH contained one neutral mutation compared to rdpAMC1, whereas the predicted RdpAMH protein had a 100% identical amino acid sequence to RdpAMC1. RdpA belonged to the class of α-ketoglutarate-dependent dioxygenases and showed a 35% identical amino acid sequence to TauD from Pseudomonas aeruginosa and percentages of identities below 33% to other members of this enzyme class (Table 3). Upstream of rdpAMH, a 1,200-bp region was found which was similar to an insertion element from Ralstonia sp. strain JS705 (Table 2). The transposase ORF comprised 1,149 nucleotides.

FIG. 2.
FIG. 2.

(A) Configuration of phenoxyalkanoic acid degradation genes in S. herbicidovorans MH; (B) comparison of the gene clusters for the modified ortho-cleavage degradation pathway in different bacteria. pJP4, R. eutropha JMP134 (M35097, U16782); pAC27, P. putida (M16964); pP51, Pseudomonas sp. (M57629). The arrows indicate the localization, size, and direction of the transcription of the genes. Dashed lines around boxes represent incompletely sequenced ORFs. Similar patterns of hatching and shading between genes indicate DNA homology or amino acid similarity of the translated gene products. The following enzymes are shown (the genes that code for them are in parentheses): α-ketoglutarate-dependent dioxygenases (rdpA and sdpA), 2,4-D transport protein (tfdK), (chloro)phenol hydroxylases (tfdB, tfdBI, and tfdBII), chlorocatechol 1,2-dioxygenases (dccAI, dccAII, tfdCI, tfdCII, clcA, and tcbC), chloromuconate cycloisomerases (tfdDI, tfdDII, clcB, and tcbD), dienelactone hydrolases (dccDI, dccDII, tfdEI, tfdEII, clcD, and tcbE), maleylacetate reductases (dccE, tfdFI, tfdFII, and tcbF), and transposases (tnpA and orfB).

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TABLE 2.

Genes and corresponding products isolated from S. herbicidovorans MH

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TABLE 3.

Identity of RdpA and SdpA to other α-ketoglutarate-dependent dioxygenases (determined by BlastP two sequences)

The cosmid 10A12 was subsequently hybridized under low-stringency conditions with a 0.7-kb EcoRI fragment of pMec10 containing the rdpAMH gene, which revealed a weak signal in addition to the rdpA-containing fragments itself (not shown). The signal could be located to a 5-kb KpnI fragment, which was cloned (pMec16) and partially sequenced. An ORF of 864 nucleotides was identified whose predicted amino acid sequence showed significant similarity to the group of α-ketoglutarate-dependent dioxygenases and highest similarity to (S)-dichlorprop/α-ketoglutarate dioxygenase from D. acidovorans MC1 (Table 2). For this reason, this ORF was designated sdpAMH. The predicted amino acid sequence for SdpAMH was 63% identical to SdpA from D. acidovorans MC1 (SdpAMC1) and 37% to TfdA from R. eutropha JMP134(pJP4) (Table 3). Upstream of sdpAMH, a similar insertion element was identified as the one in the region upstream of rdpAMH. However, the complete ORF of this element was not present on the insert of pMec16.

With the primer pair tfdBup and tfdBlow, a 1.1-kb PCR-fragment was amplified from total DNA of S. herbicidovorans MH, cloned (pMec23), and sequenced. The insert was then used to hybridize cosmids 10A12, 9H11, and 11D11, which harbor other genes of phenoxyalkanoic acid degradation. A 5.5-kb SalI fragment from 11D11 was subsequently cloned (pMec25) and partially sequenced. The sequence revealed two complete ORFs and one incomplete ORF, whose deduced amino acid sequences showed similarity to known proteins in the databank (Table 2). The deduced amino acid sequence of the incomplete ORF at the left end of the insert was similar to CadR from Bradyrhizobium sp. strain HW13 and was therefore tentatively designated cadR (25). The ORF downstream showed low similarity (<26%) to the C-terminal region of outer membrane receptor proteins. The ORF starting at nucleotide 2654 and comprising 1,800 bp showed similarity to a dichlorophenol hydroxylase gene and was designated tfdB (Table 2).

Sequence determination and organization of two chlorocatechol degradation gene clusters.To determine whether genes for chlorocatechol degradation through an ortho-cleavage pathway were present in the genome of strain MH, a PCR was carried out with the conserved primer set CCDb and CCDe (31). Amplified fragments were cloned and sequenced to determine the nature of the insert. One clone (pMec1) was found to contain a part of a putative chlorocatechol 1,2-dioxygenase gene (Table 1). The cosmid library was then hybridized with the pMec1 insert to retrieve the complete gene for chlorocatechol 1,2-dioxygenase. Interestingly, two different fragments of cosmids 9H11 and 11D11 were hybridizing, which were both subcloned and sequenced (pMec2 and pMec4). The insert of pMec4 revealed three ORFs, the deduced amino acid of the sequence of one of which had significant similarity to chlorocatechol 1,2-dioxygenases (Table 2). This ORF was tentatively designated dccAI, for dichlorocatechol 1,2-dioxygenase. The exact start of dccAI could not be determined completely, since several alternative possible start codons occurred in sequential position (i.e., at positions 265, 367, 400, 472, and 514). However, on the basis of the position of a putative Shine-Dalgarno sequence (AGGAGA at position 500), we propose the dccAI gene start at position 514. Interestingly, flanking the dccAI gene were ORFs putatively encoding a dienelactone hydrolase (designated dccDI) (Fig. 2A; Table 2) and a maleylacetate reductase. The maleylacetate reductase ORF was not completely recovered on the insert of pMec4.

The left end of the insert of pMec2 contained a part of a chlorocatechol 1,2-dioxygenase gene. To retrieve the complete ORF, a 4.5-kb BamHI fragment from cosmid 9H11, which partially overlapped the insert of pMec2, was cloned (pMec12) and sequenced. The whole ORF encoding the putative chlorocatechol 1,2-dioxygenase was obtained and tentatively designated dccAII, since it was not identical to dccAI. Flanking the dccAII on the inserts of pMec2 and pMec12 were ORFs for another dienelactone hydrolase and a putative transport facilitator protein (Table 2; Fig. 2A). However, the second possible ORF for dienelactone hydrolase (dccDII) was interrupted by a part of an insertion element identical to IS6100. The ORF upstream of dccAII most likely codes for a transporter protein similar to the TfdK 2,4-D transporter. Between dccAII and tfdK, the sequence revealed part of another insertion element (ISCc3). The sequence information thus indicated that S. herbicidovorans MH harbored at least two complete genes for a chlorocatechol 1,2-dioxygenase, which were both located upstream of a (partial) dienelactone hydrolase gene. Evidence was also found for a chloromaleylacetate reductase upstream of dccAI. Except for a gene for chloromuconate cycloisomerase, this would provide an almost complete modified ortho-cleavage pathway to strain MH.

Expression of DccAI and DccAII in E. coli BL21(DE)(pLysS).The dccAI and dccAII ORFs were expressed in E. coli BL21(DE3)(pLysS) harboring pMec9 and pMec21. In both cell extracts, bands of the expected sizes of 30.9 and 27.9 kDa, respectively, were seen on sodium dodecyl sulfate-polyacrylamide gel electrophoresis gels (not shown), which verified the translation of both genes in E. coli. Incubation of the cell extracts with 4-methylcatechol clearly showed the appearance of 4-methylmuconate with a maximal absorption at 255 nm (not shown). The substrate range for both enzymes was determined in cell extracts from E. coli BL21(DE3)(pLysS) by incubation with different substituted catechols (Table 4). DccAI and DccAII exhibited essentially the same substrate range (Table 4). Both enzymes converted chloro- and methyl-substituted catechols; the highest activity was detected with 4-methylcatechol, whereas 3,5-dichlorocatechol was preferred to monosubstituted catechols and catechol. 4,5-Dichlorocatechol was also converted, although at very low rates. No activity was measured with tetrachlorocatechol, p-nitrocatechol, protocatechuate, and hydroquinone.

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TABLE 4.

DccAI and DccAII activities with different substrates measured in cell extract of E. coli BL21(DE3)(pLysS) harboring pMec9 or pMec21

Expression of rdpA, sdpA, dccAI, and dccAII.To investigate whether rdpA, sdpA, and dccAI/II were involved in phenoxyalkanoic acid degradation in S. herbicidovorans MH, we grew the strain in batch cultures with pyruvate and pulsed the culture with (R)-, (S)-mecoprop, or 2,4-D. The formation of specific mRNAs was then determined by dot blot hybridizations (Fig. 3). mRNAs for sdpA and rdpA were always present, irrespective of the addition of mecoprop and 2,4-D, but mRNA for dccAI/II appeared specifically after exposure to both mecoprop and 2,4-D (Fig. 3). It has to be noted, however, that the hybridization experiments cannot distinguish between dccAI and dccAII because of their high similarity. These results strongly suggest that mecoprop and 2,4-D induce a modified ortho-cleavage pathway involving dccAI and/or dccAII.

FIG. 3.
FIG. 3.

Dot blot hybridization of total RNA isolated from batch cultures of S. herbicidovorans MH after induction with (R)-, (S)-mecoprop, or 2,4-D. Antisense mRNA probes used for hybridization are indicated on the left. (R)-, (S)-mecoprop and 2,4-D on the top show the induction substrate; DMSO represents the uninduced state. a, b, and c represent individual replicates.

DISCUSSION

Phenoxyalkanoic acid degradation in S. herbicidovorans MH. Here we studied the genetic background of phenoxyalkanoic acid degradation in S. herbicidovorans strain MH. Several candidate genes were isolated, which showed similarity at the amino acid level to other proteins involved in phenoxyalkanoic acid and chlorocatechol degradation. Strain MH harbors two genes encoding two α-ketoglutarate-dependent dioxygenases (named RdpAMH and SdpAMH), which were different from the archetypal TfdA enzyme (Table 3) but very similar to two dioxygenases from D. acidovorans MC1. Enzymes from D. acidovorans MC1 catalyze the enantioselective reaction of dichlorprop to dichlorophenol (55). RdpAMH and SdpAMH have been expressed in E. coli and indeed catalyze the enantioselective conversion of mecoprop and dichlorprop (T. A. Müller, unpublished data). This strongly suggests that RdpAMH and SdpAMH catalyze the first step in the degradation pathway of mecoprop and dichlorprop in strain MH (Fig. 1). Both genes were constitutively expressed and were not specifically induced upon exposure to mecoprop or 2,4-D. Previously, however, it was observed that the (R)-specific enzyme is only present when strain MH is grown on (R)-mecoprop, (R)-dichlorprop, or their racemates, whereas (S) activity is always present except when the strain is grown on the (R)-enantiomer (38). Therefore, it might be that expression of rdpAMH and sdpAMH is further regulated on the translational or posttranslational level.

S. herbicidovorans MH was shown to harbor two distinct chlorocatechol 1,2-dioxygenases genes, designated dccAI and dccAII. Both genes were induced when strain MH was exposed to mecoprop or 2,4-D. When expressed in E. coli, DccAI and DccAII showed activity toward different substituted catechols but preferred 3,5-dichlorocatechol to monosubstituted catechols. Both enzymes showed essentially the same substrate specificities. From these results, we conclude that DccAI, DccAII, or both are involved in dichlorprop, mecoprop, and 2,4-D degradation in S. herbicidovorans MH. However, we do not know whether the two dichlorocatechol 1,2-dioxygenases have different roles.

We hypothesize that the tfdB-like gene is involved in mecoprop and dichlorprop degradation in strain MH, although this was not specifically tested here. The finding of other genes from the modified ortho-cleavage pathway (i.e., dienelactone hydrolase and maleylacetate reductase) is further evidence to propose that 3,5-dichlorocatechol and 3-methyl-5-chlorocatechol are processed through a metabolic pathway similar to the one for 2,4-D degradation in R. eutropha JMP134(pJP4) (Fig. 1). Currently, no information is available concerning the location of a chloromuconate cycloisomerase gene, the product of which would be needed to complete the pathway.

Genetic organization of phenoxyalkanoic degradation.The genes for phenoxyalkanoic acid degradation in S. herbicidovorans MH were not organized in one or two operons but scattered over the genome. The rdpAMH and sdpAMH gene were located on the same cosmid but separated by at least 10 kb. The genes for chlorophenol and chlorocatechol degradation were organized in three clusters, which were located on different cosmids. Such a gene organization is in contrast to that of the tfd genes for 2,4-D degradation in R. eutropha JMP134(pJP4), which are present in one compact region of 22 kb (29). A dispersed gene organization seems to be common for Sphingomonas strains. For instance, the pcp genes coding for pentachlorophenol degradation in Sphingobium chlorophenolicum ATCC 39723 are organized on noncontiguous parts of the DNA (8). A scattered gene organization was also found for the dioxin-degradative genes in Sphingomonas sp. strain RW1 (2). The lin genes for lindane degradation in Sphingomonas paucimobilis are present in five different operons and unlinked clusters (28). This contrasting gene organization in sphingomonads may have consequences for regulation of pathway expression. Expression of rdpAMH and sdpAMH in strain MH was constitutive, whereas the dccAI/II genes were inducibly expressed. This scheme is similar to the one for expression of the lindane pathway in S. paucimobilis, where the first three genes are constitutively expressed and only the genes for chlorohydroquinone conversion are inducible (35). In contrast, the 2,4-D pathway in R. eutropha JMP134(pJP4) is very tightly regulated (32, 33). It might be that the organization observed in S. herbicidovorans is not optimized yet and that regulatory networks controlling rdpA and sdpA expression will evolve when the strain remains being exposed to phenoxyalkanoic acid-containing environments. On the other hand, the absence of inducible expression might just be a different strategy found in sphingomonads compared to pseudomonads, which nonetheless results in functional catabolic pathways. When considering that inducible catabolic pathways often seem to need a threshold concentration before induction takes place, it may even be an advantageous strategy to continuously produce the first enzymes necessary for the pathway.

Potentially two clusters might be involved in chlorocatechol degradation in strain MH. Both clusters clearly have a different gene order than the typical operons for chlorocatechol such as the clcABD genes from P. putida (pAC27) (14), the tfdCDEF genes in R. eutropha JMP134(pJP4) (9, 19, 20, 40) and the tcbCDEF genes in Pseudomonas sp. strain P51 (54) (Fig. 2B). It was noteworthy that a gene for a chloromuconate cycloisomerase was not present within the regions sequenced from strain MH. Furthermore, cluster II was flanked by sequences with clear homology to (parts of) IS elements that even interrupted dccDII; this points to (recent) genetic rearrangements.

Evolutionary aspects of the phenoxyalkanoic acid degradation pathway in S. herbicidovorans MH.It has been proposed that phenoxyalkanoic acid degradation genes are highly mobile and thus are easily transferred among bacteria (17, 24, 34, 50, 51). On the other hand, gene flow is assumed to be less frequent between Alphaproteobacteria and Beta- and Gammaproteobacteria than among Beta- and Gammaproteobacteria (17). The unique combination and organization of the phenoxyalkanoic acid degradation genes in S. herbicidovorans MH indicate that a recent gene exchange between Alpha- and Betaproteobacteria must have taken place. This can be concluded from the finding that RdpAMH is 100% identical to RdpAMC1 from D. acidovorans MC1, a member of the Betaproteobacteria, and also clearly contradicts the hypothesis that tfdA-like genes of Alphaproteobacteria and Beta- and Gammaproteobacteria have arisen by divergent evolution (23). The transposase genes located next to rdpAMH and sdpAMH also support the hypothesis of gene mobilization. The transposase gene located nearby rdpAMH is very similar to a presumably promiscuous IS element which was implicated in transferring the toluene dioxygenase from Ralstonia sp. strain JS745 to Ralstonia sp. strain JS705 (37).

In contrast to the α-ketoglutarate-dependent dioxygenase genes, chlorocatechol 1,2-dioxygenases from Beta- and Gammaproteobacteria present in the database are less related to DccAI/II than they are among themselves. In a phylogenetic analysis, strain MH's chlorocatechol 1,2-dioxygenases clustered together with chlorocatechol 1,2-dioxygenases from gram-positive bacteria forming a separate branch (not shown), which is in agreement with the results obtained by Eulberg et al. (12). Six different groups of chlorocatechol 1,2-dioxygenase could be defined: (i) ClpC, (ii) Sphingomonas homologs, (iii) TfdC_pJP4 homologs, (iv) the TcbC-TetC group, (v) TfdCII_pJP4 homologs, and (vi) the ClcA homologs. Subgroups i and ii build new branches, whereas the classification of subgroups iii to vi agrees with the one described by Armengaud et al. (3). The two novel groups comprise sequences from Alphaproteobacteria and support our hypothesis that Sphingomonas strains have not exchanged their chlorocatechol 1,2-dioxygenase genes recently with those from Beta- and Gammaproteobacteria. The phenoxyalkanoic acid degradation pathway of strain MH has been demonstrated to be unique in its composition, genetic organization, and regulation, and it will be important to further study its characteristics in order to broaden our understanding of pathways different from the canonical 2,4-D degradation pathways in Beta- and Gammaproteobacteria.

ACKNOWLEDGMENTS

T.A.M. was supported by Swiss National Science Foundation grant NF-3100-055468.

FOOTNOTES

    • Received 2 March 2004.
    • Accepted 22 June 2004.
  • ↵*Corresponding author. Mailing address: Department of Fundamental Microbiology, BÂtiment de Biologie, University of Lausanne, CH-1015 Lausanne, Switzerland. Phone: 41 21 692 5630. Fax: 41 21 692 5635. E-mail: JanRoelof.VanDerMeer{at}imf.unil.ch.

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