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Applied and Environmental Microbiology, November 2008, p. 6631-6638, Vol. 74, No. 21
0099-2240/08/$08.00+0     doi:10.1128/AEM.01192-08
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Involvement of a Novel Enzyme, MdpA, in Methyl tert-Butyl Ether Degradation in Methylibium petroleiphilum PM1

Radomir Schmidt,^1* Vince Battaglia,¹ Kate Scow,¹ Staci Kane,² and Krassimira R. Hristova¹

Department of Land, Air and Water Resources, University of California, Davis, California,¹ Lawrence Livermore National Laboratory, Livermore, California²

Received 28 May 2008/ Accepted 7 September 2008

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Methylibium petroleiphilum PM1 is a well-characterized environmental strain capable of complete metabolism of the fuel oxygenate methyl tert-butyl ether (MTBE). Using a molecular genetic system which we established to study MTBE metabolism by PM1, we demonstrated that the enzyme MdpA is involved in MTBE removal, based on insertional inactivation and complementation studies. MdpA is constitutively expressed at low levels but is strongly induced by MTBE. MdpA is also involved in the regulation of tert-butyl alcohol (TBA) removal under certain conditions but is not directly responsible for TBA degradation. Phylogenetic comparison of MdpA to related enzymes indicates close homology to the short-chain hydrolyzing alkane hydroxylases (AH1), a group that appears to be a distinct subfamily of the AHs. The unique, substrate-size-determining residue Thr⁵⁹ distinguishes MdpA from the AH1 subfamily as well as from AlkB enzymes linked to MTBE degradation in Mycobacterium austroafricanum.

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Use of methyl tert-butyl ether (MTBE) as a fuel oxygenate over the last 2 decades has resulted in contamination of numerous subsurface drinking water supplies. The oxygenate is now banned in California and is being phased out elsewhere in the United States (5). At present, little is known concerning the biochemistry and genetics of aerobic MTBE metabolism, which involves a novel ether cleavage reaction described primarily for cometabolic MTBE-degrading organisms (29, 33). Physiological studies have suggested that enzymes involved in the initial step of the MTBE biodegradation pathway belong to the alkane hydroxylase (AH) family (21, 29, 33). In Methylibium petroleiphilum PM1, an enzyme closely related to AHs (MdpA) has been the prime candidate for the enzyme involved in the first step of MTBE degradation (13, 17).

M. petroleiphilum PM1 is a methylotroph representing a new species within the Rubrivivax group (Comamonadaceae family) of the beta subclass of the Proteobacteria (23). Strain PM1 was isolated from a sewage treatment plant biofilter that was used for treating discharge from oil refineries (10) and is one of few pure culture isolates that can completely degrade the fuel additive MTBE (2, 10, 23). Pilot and field studies have demonstrated the efficacy of aerobic bioremediation of MTBE by PM1 (3, 5a, 27, 32, 36). Furthermore, PM1-like bacteria (98 to 99% similar based on 16S rRNA gene sequences) have been shown to be naturally occurring in a number of MTBE-contaminated aquifers in California (12, 16, 18). In situ studies correlating total and PM1-like bacterial cell counts with MTBE degradation rates suggest that PM1-like organisms play a significant role in MTBE biodegradation under aerobic conditions in California aquifers (12).

The whole-genome sequence of PM1 was obtained to provide a framework for examining MTBE degradation pathways and other important metabolic pathways in this bacterium (17). A subsequent microarray study examined the changes in gene expression levels in PM1 grown with either ethanol or MTBE as the sole source of carbon. Genetic and expression analyses revealed a 10-kb region of the PM1 megaplasmid that carries all three components necessary for the production of a functional AH system (monooxygenase, rubredoxin, and rubredoxin reductase). Individual genes of the predicted MTBE degradation pathway showed 1.5- to 13-fold upregulation in cultures grown in the presence of MTBE compared to growth on ethanol (13).

The choice of genetic techniques for working with M. petroleiphilum PM1 is limited, because the organism is naturally resistant to a wide spectrum of antibiotics, and it readily forms spontaneous mutants against a number of antibiotics. In addition, it does not express resistance to at least two antibiotics it is naturally sensitive to (ampicillin and tetracycline) when the resistance genes are provided in trans. In this study, we developed a genetic system for M. petroleiphilum PM1, building on previous results using random mutagenesis based on the pTnMod-SmO vector (4, 17). Approaches we used included efficient electroporation of PM1, targeted mutagenesis based on the Epicentre in vitro mutagenesis system (7), complementation using pBBR1MCS-2 based vectors (19), and establishment of a useful DNA fragment limit for homologous recombination in PM1. In this report, we show how inactivation of mdpA demonstrates that the protein product (MdpA) is responsible for MTBE removal, while it does not play a direct role in tert-butyl alcohol (TBA) degradation. The results of our complementation and inhibitor studies confirm that MdpA is the sole enzyme responsible for MTBE degradation in PM1. Functional and phylogenic studies of MdpA indicate that it is a unique enzyme closely related to short- to medium-chain AHs.

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Bacterial strains and culture conditions.
M. petroleiphilum PM1 cultures were routinely grown in 1/3x tryptic soy broth (TSB) at 28°C with rotary shaking at 150 rpm or on 1/3x TSB agar at 28°C. When required, antibiotics were added at the following final concentrations: kanamycin (Km), 50 µg/ml; streptomycin (Sm), 50 µg/ml; spectinomycin (Spm), 50 µg/ml. For carbon source utilization and degradation experiments, M. petroleiphilum PM1 was grown in mineral salts medium (MSM; Tris-HCl, 0.13 M; KH₂PO₄, 0.023 M; K₂HPO₄, 0.025 M; CaCl₂, 0.027 M; NaHCO₃, 0.2 M; MgSO₄, 0.05 M; EDTA, 0.0288 mM; and NH₄Cl, 0.27 M) supplemented with trace elements (CoCl₂, 0.25 µM; CuSO₄, 0.3 µM; FeCl₃, 40 µM; H₃BO₃, 50 µM; MnCl₂, 10 µM; Na₂MoO₄, 0.1 µM; ZnSO₄, 0.8 µM). Carbon sources included MTBE (250 mg/liter), ethanol (790 mg/liter), sodium acetate (1 g/liter), TBA (250 mg/liter), and n-pentane (250 mg/liter). PM1 is capable of growth on MSM with up to 1,000 mg/liter MTBE or up to 7.9 g/liter ethanol. Cells were grown at 28°C in 50-ml batch cultures in 150-ml glass bottles with rotary shaking at 150 rpm. Escherichia coli DH5 cells were used for all transformations that involved vectors carrying Sm resistance. For all other transformations, E. coli TOP10 (Invitrogen) cells were used. All E. coli cultures were grown on Luria-Bertani (LB) agar at 37°C. All strains and plasmids used in this study are listed in Table 1.

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TABLE 1. Bacterial strains, plasmids, and transposons used in this study

Construction of transposon vector for disruption of mdpA.
In order to perform insertional inactivation of mdpA, a custom EZ-Tn5<SmQ> transposon was constructed. This in vitro mutagenesis vector with Sm^r Spm^r was based on the pMOD-2<MCS> vector (Epicentre) with the addition of the Sm resistance cassette from the pTnMod-SmO vector (4). Briefly, PCR primers were designed to include the aadA3 gene and its promoter (P2) from pTnMod-SmO while excluding the integrase gene upstream of the aadA3 gene. The PCR product was cloned into pCR2.1 TOPO (Invitrogen). The resulting plasmid, containing aadA3, was digested with EcoRI and cloned into the EcoRI site in the multiple cloning site (MCS) of pMOD-2<MCS>, resulting in EZ-Tn5<SmQ>. Following the standard Epicentre in vitro mutagenesis protocol for generation of custom transposons, EZ-Tn5<SmQ> was amplified by PCR using the pMOD<MCS> forward and reverse PCR primers (Epicentre) and purified with a QIAquick PCR purification kit (Qiagen).

Construction of pK18-mdpA and mdpA knockout strains.
Five sets of PCR primers were designed to amplify incrementally larger regions surrounding the mdpA gene in PM1 (Table 2). For the determination of homologous-region size requirement, five constructs of different size flanking the same EZ-Tn5<SmQ> insert were constructed. Primer set 5 was used to amplify a 4.5-kb region surrounding mdpA. The PCR product was cloned into pCR2.1 TOPO (Invitrogen). A correct clone was selected and checked by sequencing, digested with HindIII/NsiI, and cloned into the HindIII/PstI sites in the MCS of pK18 (24). The resulting plasmid, pK18-mdpA5, determined by restriction digestion, was selected for in vitro mutagenesis. Equimolar amounts (0.04 pmol) of pK18-mdpA5 and EZ-Tn5<SmQ> were mixed together with the transposase and incubated for 2 h at 37°C. The reaction was stopped and the DNA was transformed into E. coli DH5 cells. Transformants were selected on LB agar containing 50 µg/ml Km and 50 µg/ml Sm. Transposon inserts were checked by PCR using primer set 1 to ensure insertion in mdpA. The exact site of insertion of likely candidates was determined by sequencing amplicons using the pMOD<MCS> forward and reverse sequencing primers (Epicentre). Primer sets 1 to 5 were used to amplify five fragments of different sizes flanking mdpA::Sm^r, the insert in pK18-mdpA5<SmQ>. The PCR products were purified and concentrated with a QIAquick PCR purification kit (Qiagen). Equimolar concentrations (0.6 pmol) of all five PCR products were individually transformed into PM1 by electroporating 50 µl of cells washed in 10% glycerol in a Bio-Rad Gene Pulser electroporator set at 1.8 kV, 200 , and 25 µF. Transformants were selected on 1/3x TSB agar containing 50 µg/ml Sm and 50 µg/ml Spm to eliminate the possibility of spontaneous mutants. Triplicate plate counts showed that electroporation carried out with the 2.1, 3.3 and 4.5 kb based vectors resulted in 20 to 30 CFU/plate, while the shorter length fragments resulted in few to no colonies. Therefore, the minimum fragment length of host DNA required for efficient homologous recombination based mutagenesis in PM1 is 2 kb.

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TABLE 2. Primers

The procedure for the mdpA insert mutant generation was essentially as described above, except that primer set 3 (Table 2) was used to amplify and clone a 2.1-kb mdpA-containing fragment from PM1 (Fig. 1) and EZ-Tn5<SmQ>#2 insert (carried on pK18-mdpA3<SmQ>) was used to inactivate the wild-type mdpA gene in PM1. Twenty Sm-resistant subclones were tested for insert presence by PCR using primer set 5 (Table 2). Subclones were tested for correct, stable insertion by PCR. An increase in fragment size from 4.5 kb to 6 kb in all 20 subclones tested, corresponding to the expected increase based on the size of the EZ-Tn5<SmQ> insert, showed that the disruption had occurred in the correct location. Further testing following growth without antibiotic selection on both defined (MSM) and rich media (1/3x TSB) showed that the <SmQ> cassette insert was stably integrated into the megaplasmid, with no loss of antibiotic resistance after >20 generations. One PCR-confirmed positive strain MP0024 (mdpA) was used for further study.

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FIG. 1. mdpA and neighboring genes. The Smr transposon inserts in mdpA are indicated. The position of the potential promoter upstream of mdpA is shown. Primer sets 1 to 5 (Table 2) are shown. EZ-Tn5<SmQ> insertion points for mdpA inactivation (#2) and recombination size requirement (#22) experiments are shown. The mdpA coding sequence is 1,200 bp, and the predicted molecular mass of the 400-amino-acid MdpA protein is 45.2 kDa.

Resting-cell experiments.
The mdpA strain MP0024 and the wild type, MP0002, were tested for growth on MSM with 0.1% (vol/vol) ethanol, 0.1% (wt/vol) sodium acetate, 250 mg/liter MTBE, or 250 mg/liter TBA as the sole carbon source. Strains MP0002 and MP0024 were tested in a resting-cell experiment for their ability to degrade MTBE and TBA. Fifty-ml cultures were grown overnight on MSM with 0.1% (vol/vol) ethanol to an optical density at 595 nm (OD₅₉₅) of 0.2, Fifty ml of each was used to inoculate 450 ml of MSM with 0.1% (vol/vol) ethanol, and the cultures were grown to early log phase (OD₅₉₅ = 0.2 to 0.4). The cells were washed twice and resuspended to a final volume of 14 ml. The final cell suspension was used to inoculate 45-ml glass microcosm bottles containing 20 ml MSM with 5 mg/liter MTBE or 5 mg/liter TBA to a final concentration equivalent to an OD₅₉₅ of 0.6. Screw caps with Mininert valves (Pierce Biotechnology, Rockford, IL) were used to seal the microcosm bottles.

Samples (1 ml) were extracted from each microcosm in a sterile laminar flow hood. Samples were stored in 10-ml-headspace vials preserved with sodium phosphate tribasic dodecahydrate at a concentration of 1% by weight and sealed with 20-mm Teflon-lined septa and aluminum crimp caps. Samples were stored no longer than a week at 4°C prior to analysis. MTBE, TBA, and ethanol were quantified on an Agilent 6890N gas chromatograph equipped with a flame ionization detector and an HP 7694 headspace autosampler. Organic compounds were separated using an Agilent HP1 capillary column (60 m by 1 µm by 0.320 µm). The output was analyzed using ChemStation revision A.10.02 software (Agilent, Santa Clara, CA).

mdpA complementation.
Primer set 4 (Table 2) was used to amplify a 3.3-kb region surrounding mdpA. The PCR product was cloned into pCR2.1 TOPO (Invitrogen). The entire EcoRI cassette was transferred to pK18, yielding pK18-mdpA4. A resulting clone was checked for the correct insert by sequencing and was subsequently digested with XbaI, and the mdpA fragment was cloned into the unique XbaI site of pUC19 (the pUC19 cloning step was carried out to simplify vector selection, since both pK18 and pBBR1MCS-2 carry Km resistance). The 1.8-kb XbaI mdpA cassette was transferred to the unique XbaI site of pBBR1MCS-2, the construct checked by sequencing and a plasmid with the correct insert selected (pBR21). Both pBR21 and pBR27 were individually transformed into PM1 by electroporation. Transformants were selected on 1/10x TSB agar containing 50 µg/ml Km. Plasmid DNA was extracted from antibiotic-resistant strains to verify transformant status. Confirmed positive strains MP0053 (pBR21) and MP0054 (pBR27) as well as a strain containing plasmid pBBR1MCS-2 (MP0051) as a negative control were used for further study.

Monooxygenase inhibitor studies.
The wild-type strain MP0002 was tested in resting-cell experiments for its ability to degrade MTBE, TBA, or ethanol in the presence of monooxygenase inhibitors. Strain MP0002 was grown in 50-ml cultures consisting of MSM with MTBE (250 mg/liter) or TBA (250 mg/liter). Fifty-ml cultures were transferred twice in MSM with the respective carbon source. Early-log-phase cultures were used to inoculate the final cultures with each carbon source. Cells were harvested in early log phase (OD₅₉₅ = 0.2 to 0.4). Cells were washed twice and resuspended in MSM, and the final cell suspension was used to inoculate 45-ml microcosm bottles containing 20 ml MSM with 50 mg/liter MTBE or 50 mg/liter TBA. When required, monooxygenase inhibitors were added to the following concentrations: acetylene, 40% vol/vol (in headspace); methimazole, 10 mM.

Sequence analysis and generation of phylogenetic trees.
Homologs of translated M. petroleiphilum PM1 coding sequences were identified using BLASTP searches against the nonredundant GenBank database from the National Center for Biotechnology Information (NCBI). Sequences were aligned and alignments were refined using ClustalW along with manual adjustments, and the Protdist program and the Neighbor program of the BioEdit package (8) were used to generate phylogenetic trees.

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MdpA structure.
MdpA contains six conserved transmembrane (TM) domains of the AH family (residues 25 to 66, 94 to 139, and 234 to 276). The three clusters of conserved histidine residues found in hydrocarbon oxygenases and desaturases (His^{142, 146}, His^{172, 176, 177}, and His^{317, 320, 321}) are also present, as is the conserved motif ²⁷⁴DYIEHYGL²⁸¹ [corresponding to NYXEHYG(L/M), found in AHs (1)]. A single conserved amino acid position apparently divides AHs into two major groups. In group AH1, an amino acid in a position facing the central channel of the integral membrane protein is a bulky side chain (either tyrosine or tryptophan), and the hydroxylase cannot degrade long-chain alkanes (35). In group AH2, the residue has a small side chain (isoleucine, leucine, or valine), and the hydroxylase degrades long-chain alkanes, apparently because they are not restricted from entering further into the AH central channel (35). An alignment of MdpA with several top BLAST hit AHs and previously characterized proteins shows that the residue Thr⁵⁹ does not belong to either of the above classes (Fig. 2). A phylogenetic tree of AHs (Fig. 3) shows clustering among the AH1 group and clear separation from the AH2 group. A phylogenetic tree of 199 full AH sequences available in the NCBI database (as of May 2008; data not shown) also showed tight clustering among putative AH1 enzymes, but only as a single, distinct branch among many on the whole tree. This finding is consistent with a prediction that short-chain AHs form a phylogenetically distinct subfamily of AHs, and the Tyr/Trp residue is only one of the distinguishing features of this group.

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FIG. 2. Sequence alignment of TM helix 2 and comparison of properties. The predicted AH grouping reflects the preferred alkane substrate (AH1, medium-chain-length alkanes; AH2, long-chain alkanes); proteins of known alkane length preference are indicated with a superscript letter "a" (34, 35). The fourth column shows the alignment of TM helix 2, with the position of the first and last residues indicated above (PM1 MdpA position). TM helix 2 runs from the periplasm to the cytoplasm. Position 59 of PM1 MdpA (equivalent to position 55 of P. putida GPo1 AlkB [35]) is indicated by an X, and the AH residue proposed by van Beilen and coworkers (35) to determine alkane chain length preference is shown in bold.

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FIG. 3. Phylogenetic tree of M. petroleiphilum MdpA against related AHs. The predicted AH1 group forms a distinctive lineage. MdpA clusters more closely with the AH1 group of AHs.

mdpA and neighboring gene structure.
The organization of genes involved in alkane oxidation varies widely among the alkane-degrading bacteria. Genes involved in alkane degradation can be distributed over the genome (31). The rubredoxin reductase genes are usually not located close to an AH gene, perhaps because they are also involved in other pathways and require independent regulation (31). In contrast, most rubredoxin genes are located immediately downstream of the AH genes. The Gordonia sp. strain TF6 AH cluster is the simplest arrangement of required genes (31). The mdpA, rubredoxin, and rubredoxin reductase genes of PM1, which are homologs of alkB, alkF/G, and alkT, respectively, map to a single 10-kb locus on the megaplasmid. Also within this locus is the putative regulator mdpC as well as a transposase gene (MpeB0605), a putative esterase gene (MpeB0604), and three hypothetical genes (MpeB0598, MpeB599, and MpeB600).

Inhibitor studies.
Methimazole is an irreversible inhibitor of flavin cofactor (FAD)-containing monooxygenases, and acetylene is a mechanism-based inhibitor or inactivator of specific monooxygenases, including alkane monooxygenase. We tested methimazole inhibition of MTBE or TBA degradation by resting cells of PM1 grown on either carbon source (Fig. 4). In general, methimazole inhibited MTBE degradation irrespective of which carbon source was utilized for cell growth (Fig. 4). Conversely, while TBA removal was inhibited by methimazole for PM1 grown on MTBE, TBA removal was unaffected by methimazole when PM1 was grown on TBA. These results suggest that a flavin-dependent monooxygenase is directly responsible for MTBE degradation but is involved only at the TBA oxidation (13) step via a regulatory mechanism, since the inhibition of the flavin monooxygenase leads to the prevention of TBA removal only in MTBE-grown cells. We also tested acetylene inhibition of MTBE degradation by resting cells of PM1 grown on MTBE. No effect of acetylene on MTBE removal was detected (data not shown). Inhibition of the first ether scission reaction in MTBE degradation by methimazole suggests that MTBE monooxygenase is a flavin-dependent enzyme and distinct from other alkane monooxygenases.

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FIG. 4. Effect of methimazole on MTBE (a) or TBA (b) removal. Wild-type PM1 cells grown in MSM with either MTBE (squares) or TBA (triangles) as the sole carbon source were resuspended in MSM, and MTBE or TBA removal was measured in the absence (filled symbols) or presence (open symbols) of 10 µg/ml methimazole. Experiments were carried out in triplicate; error bars show 1 standard deviation. Variation in apparent initial concentration is due to the volatile nature of MTBE and TBA.

MTBE degradation inducibility.
We compared MTBE degradation by PM1 resting cells grown in the presence of various carbon sources. We performed resting-cell experiments to specifically observe the metabolism of MTBE independent of growth by the organism. PM1 cells were grown to mid-log phase in the presence of either 500 µg/ml MTBE, 773 µg/ml ethanol, or 1,000 µg/ml sodium acetate, washed, and resuspended in MSM plus 5 µg/ml MTBE. Previous results predicted a higher MTBE degradation rate with MTBE-grown resting cells, and therefore these cells were also tested in the presence of 50 µg/ml MTBE. In addition, ethanol-grown cells were assayed in the absence and presence of the translational inhibitor chloramphenicol (Chl). MTBE removal in the presence of Chl indicates a low level of constitutive expression of MdpA in PM1 grown on ethanol (Fig. 5; Table 3. Degradation proceeded most) rapidly in the MTBE-grown cells, with only trace amounts of MTBE (<0.05 mg/liter) detected in the first samples (30 min for 5 µg/ml; 1 h for 50 µg/ml). The degradation rate is therefore at least 14.8 nmol/min/mg protein with 5 µg/ml MTBE and at least 72.6 nmol/min/mg protein with 50 µg/ml MTBE. The six- to eightfold-lower rate of MTBE removal in the presence of Chl (Table 3) and the rapid MTBE degradation in MTBE grown cells show that MdpA is strongly induced by MTBE.

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FIG. 5. MTBE removal by wild-type PM1 cultures grown on MTBE (open circles), ethanol (filled diamonds), and ethanol and by resting cells incubated with Chl (filled circles). The sterile control is indicated by filled squares. Note that MTBE-grown cells were also incubated in the presence of either 10 µg/ml or 50 µg/ml MTBE, and in both cases only trace amounts of MTBE (>10 ng/ml) were detected in the samples taken at 1 h.

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TABLE 3. Specific degradation rates of wild-type PM1 and mutants on oxygenatesa

mdpA knockout; MTBE and TBA degradation.
To test the predicted function of MdpA in MTBE degradation, we disrupted the mdpA gene in vitro with EZ-Tn5<SmQ>#2 (Fig. 1) and then replaced the wild-type copy of mdpA in PM1 with the mutant version by homologous recombination. We compared the specific MTBE and TBA removal capacity of the mdpA mutant (MP0024) to that of the M. petroleiphilum PM1 wild-type strain MP0002. In a resting-cell experiment performed with cultures grown on ethanol, the wild type and MP0024 were able to degrade TBA at equivalent rates, but only the wild type was able to metabolize MTBE (Fig. 6). These results are the first direct linkage of an alkane monooxygenase family protein to the initial reaction of MTBE oxidation. MP0024 grew on TBA but not on MTBE as the single carbon source. The results are also consistent with the results of the inhibitor studies that predicted distinct, unique enzymes for the degradation of MTBE and TBA. Moreover, as no reduction in the observed rate of TBA removal occurred for the mdpA mutant (Fig. 6) it seems likely that MdpA does not contribute to TBA degradation in PM1.

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FIG. 6. Comparison of wild-type MP0002 (diamonds) and mdpA mutant MP0024 (squares) removal of MTBE (filled symbols) and TBA (empty symbols) by resting cells of M. petroleiphilum PM1 grown on ethanol. Experiments were carried out in triplicate; error bars show 1 standard deviation. Variation in apparent initial concentration is due to the volatile nature of MTBE and TBA.

mdpA complementation.
When initial experiments with IncP-based broad-host-range vectors for replication in PM1 failed, other vectors were tried. Although the pBBR-based vector pBBR1MCS-2 was also quickly eliminated in cultures not under antibiotic selection (5 to 10 generations), the plasmid was recoverable even after several transfers in various media with 100 µg/ml Km. An XbaI fragment containing mdpA plus the presumed promoter (221 bp upstream) was cloned into pBBR1MCS-2. The wild type (MP0002), the mdpA mutant strain carrying the pBBR1MCS-2 vector alone (MP0051), and the mdpA mutant strain carrying the complementation plasmid (MP0053) were used for further experiments.

The complementation strain did not grow on MTBE. However, a resting-cell experiment carried out with MP0002, MP0051, and MP0053 grown on ethanol showed a similar pattern of MTBE degradation for the wild-type (MP0002) and mdpA⁺ (MP0053) strains (Fig. 7a; Table 3). No MTBE removal was observed in the presence of the pBBR1MCS-2 vector alone (strain MP0051). The fate of TBA as the substrate was also monitored in resting-cell experiments. Similar conversion rates were observed for all three strains for TBA degradation (Fig. 7b; Table 3).

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FIG. 7. Complementation studies of mdpA. The wild type (diamonds), the mdpA mutant carrying the pBBR2MCS-2 (squares), and the mdpA mutant carrying the complementation plasmid pBR21 (triangles) were tested for removal of MTBE (a) or TBA (b) by resting cells. Experiments were carried out in triplicate; error bars represent 1 standard deviation. Variation in apparent initial concentration is due to the volatile nature of MTBE and TBA.

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Despite previous research on AlkB-type AHs, the details of the mechanism of action of these membrane-bound enzymes remain elusive due to the difficulty of obtaining crystallography data (26). On the other hand, it has been established that the six TM domains of the enzyme form a deep cleft; alkanes of various lengths can penetrate into this cleft and are then oxidized.

Recently, bulky and small side chains at a unique conserved amino acid position were shown to play a major role in determining whether AHs are active on short- to medium-chain (AH1) or long-chain (AH2) alkanes (35). A phylogenetic tree of AHs (Fig. 3) shows separation of AH1 from AH2 enzymes. Single residue differences are not likely to produce homogeneous grouping, so the AH1 cluster is likely to share evolutionary lineage and therefore features other than the key Tyr or Trp residues.

An alignment of MdpA with previously characterized proteins shows that the key residue Thr⁵⁹ does not belong to either the AH1 or AH2 class (Fig. 2). Because Thr⁵⁹ is likely to play a role in substrate range determination in MdpA, we searched for other proteins with the same residue at this position. A thorough search of all 199 complete alkane-1-monooxygenases in the NCBI database (as of May 2008) found only one other branch that contained proteins with Thr as the determinative residue, a tight cluster of putative AHs from members of the family Rhodobacteraceae. These proteins showed only 30% identity to MdpA on average, were not associated with a known substrate range, and did not cluster with the AH1 proteins (data not shown).

We compared MdpA to other enzymes reported to be active in MTBE degradation. Only partial sequences of predicted MTBE monooxygenases from Mycobacterium austroafricanum strains were available (22), but the truncated sequences were identical to each other and differed in only one residue in a 114-residue overlap with Mycobacterium vanbaalenii PYR-1 AlkB. The M. vanbaalenii PYR-1 enzyme therefore provided a reasonable estimate of the position of these enzymes within the phylogeny. All mycobacterial AlkBs, including M. vanbaalenii PYR-1, clustered closely within AH2 (Fig. 3). MdpA shows 46 to 50% identity to the AH1 proteins, slightly higher than the 42% identity it shares with M. vanbaalenii PYR-1 AlkB. This is consistent with the deep branching of MdpA and may indicate that MdpA represents a unique subfamily of AHs.

We have established an efficient approach for genetic manipulation of PM1, including a definition of a useful DNA fragment limit for homologous recombination in PM1. Based on the insertional inactivation and complementation results, we demonstrated that MdpA, the predicted protein product of mdpA, is essential for MTBE degradation in strain PM1 (Fig. 6 and 7). We recently showed by reverse transcriptase quantitative PCR that mdpA expression is induced on MTBE (13). The induction of MTBE removal observed in this study (Fig. 5; Table 3) is consistent with this result. We are currently investigating whether lack of growth on MTBE by the complementation strain is due to a polar effect of the original insertion or to an effect of the expression of mdpA from the complementation plasmid.

A possible MdpA mode of action is hydroxylation of the methyl ether group of MTBE, a mechanism predicted for other AlkB enzymes (21, 29, 33). AH AlkB of Pseudomonas putida GPo1 degrades MTBE with low efficiency following growth on n-octane (28). The involvement of AH in cometabolic degradation of MTBE is also predicted in Pseudomonas mendocina KR-1 (30) and M. austroafricanum ATCC 29678 (formerly Mycobacterium vaccae JOB5) (29). AlkB has been linked to both MTBE and TBA degradation in M. austroafricanum IFP 2012 (22).

The MTBE hydrolysis rate for MTBE-grown PM1 (72.6 nmol/min/mg protein) is more rapid than that reported for Hydrogenophaga flava ENV735 grown on MTBE (46 nmol/min/mg protein) (11). There was no reduction in the observed rate of TBA degradation by the mdpA mutant (Fig. 6; Table 3) demonstrating that MdpA does not play a role in TBA removal in PM1. This is in contrast to the prediction that AlkB is responsible for both MTBE and TBA removal in M. austroafricanum strains (22).

We tested the effect of two potential MdpA inhibitors. Acetylene is a mechanism-based inactivator of several monooxygenases, including methane, ammonia, and butane monooxygenases (9, 14, 25). Acetylene has been reported as an inhibitor of monooxygenase activity linked to MTBE degradation in Arthrobacter strain ATCC 27778 (20) and to both MTBE and TBA removal in M. vaccae JOB5 (15, 30) and M. austroafricanum IFP2012 (6). MTBE degradation in PM1 was not inhibited by acetylene (data not shown), similar to the reported lack of inhibition of MTBE removal in H. flava ENV735 (11). Further work is required to determine if acetylene can be used to differentiate functionally distinct AHs involved in MTBE removal. The results of the methimazole inhibition study are consistent with a flavin-dependent monooxygenase, such as MdpA, being responsible for MTBE degradation (Fig. 4). Interestingly, we observed that methimazole reduced TBA hydrolysis in PM1, but only in cultures initially grown with MTBE as the sole carbon source (Fig. 4). This is consistent with the involvement of a MTBE monooxygenase at the TBA hydrolysis step via a regulatory mechanism, rather than directly. TBA induction of a second enzyme system capable of transforming TBA but not MTBE is unlikely, since we did not observe induction of TBA removal activity when MTBE-grown cells were washed and resuspended in TBA MSM (with methimazole), even after a significant lag period.

The genetic system developed in this study confirmed that mdpA is essential for MTBE degradation in PM1. Current work is being focused on disruption and complementation of other genes predicted to be involved in TBA oxidation and other downstream catabolic reactions, as well as the expression, purification, and characterization of MTBE pathway enzymes identified by inactivation. The genetic system described in this study is a useful tool for expanding our understanding of the novel catabolic steps involved in aerobic MTBE degradation in PM1 and genetically related MTBE-degrading microorganisms.

 
This publication was made possible by grant 5 P42 ES004699 from the National Institute of Environmental Health Sciences (NIEHS), NIH.

The contents are solely the responsibility of the authors and do not necessarily represent the official views of the NIEHS, NIH.

Lawrence Livermore National Laboratory is operated by Lawrence Livermore National Security, LLC, for the U.S. Department of Energy, National Nuclear Security Administration, under contract DE-AC52-07NA27344.

 
* Corresponding author. Mailing address: Department of Land, Air and Water Resources, University of California, Davis, One Shields Ave., Davis, CA 95616. Phone: (530) 752-1488. Fax: (530) 752-1552. E-mail: radschmidt{at}ucdavis.edu

Published ahead of print on 12 September 2008.

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Applied and Environmental Microbiology, November 2008, p. 6631-6638, Vol. 74, No. 21
0099-2240/08/$08.00+0     doi:10.1128/AEM.01192-08
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