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1 Institut Français du Pétrole, 1–4, avenue de Bois-Préau, F-92852 Rueil-Malmaison, France
2 Biotechnology Research Institute, NRCC, 6100 Royalmount Avenue, Montreal, Quebec, Canada H4P2R2
Correspondence
Françoise Fayolle-Guichard
francoise.fayolle{at}ifp.fr
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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The GenBank/EMBL/DDBJ accession number for the genomic DNA sequence reported in this paper is DQ147773.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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The widespread use of MTBE has resulted in accidental releases from leaking storage tanks and spills into groundwater and surface water (Johnson et al., 2000
; US Environmental Protection Agency, 1999). The persistence of MTBE in aquifers is attributed to its high aqueous solubility and its recalcitrance to biodegradation under aerobic and anaerobic conditions. Because of the possible toxic effect of MTBE, the US Environmental Protection Agency (1999) has classified this compound as a potential carcinogen and set the maximal threshold of MTBE allowed in drinking water at 20–40 µg l–1 (US Environmental Protection Agency, 1997). The environmental and health concerns related to MTBE have led to a progressive phase-out in the US. However, there is a need for remediation technologies when aquifers are impacted by MTBE, but the usual physico-chemical techniques are generally neither efficient nor cost-effective.
The biodegradation potential of MTBE has been investigated and a few strains, such as Rubrivivax gelatinosus PM1 (Deeb et al., 2001; Hanson et al., 1999), Hydrogenophaga flava ENV735 (Hatzinger et al., 2001) and Mycobacterium austroafricanum IFP 2012 (François et al., 2002), have been isolated. More recently, other strains able to grow in the presence of MTBE have been isolated from different environments: strain UC1, closely related to R. gelatinosus PM1, and UC3, closely related to M. austroafricanum IFP 2012, were isolated from an MTBE-degrading biofilter (Pruden & Suidan, 2004); strain L108 was isolated from an MTBE-contaminated aquifer in Germany (Rohwerder et al., 2004); and M. austroafricanum IFP 2015 was isolated from the drain water of an MTBE-supplemented gasoline storage tank (Lopes Ferreira et al., 2005). The MTBE biodegradation pathway of M. austroafricanum IFP 2012 has been partially characterized by identifying several degradation intermediates, such as tert-butyl alcohol (TBA), which frequently accumulates in aquifers contaminated by MTBE (François et al., 2002). We recently detected 2-methyl 1,2-propanediol (2-M1,2-PD) as the product of TBA oxidation in M. austroafricanum IFP 2012 and IFP 2015 (Lopes Ferreira et al., 2005), and demonstrated its transformation to 2-hydroxyisobutyric acid (HIBA) in resting-cell experiments. 2-Methyl 1,2-propanediol (2-M1,2-PD) has been reported by Steffan et al. (1997), using propane-oxidizing bacteria able to degrade MTBE by cometabolism, and by Smith et al. (2003), using Mycobacterium vaccae JOB5 as the product of TBA degradation.
Using individual bacterial isolates, growth in the presence of MTBE is generally slow and the cellular yield is poor (Fayolle & Monot, 2005; Fortin et al., 2001). Several possible limitations to growth in the presence of MTBE have been suggested by Salanitro (1995). In the case of M. austroafricanum IFP 2012, the crucial role of cobalt divalent cations in the biodegradation of HIBA, one of the MTBE degradation intermediates, has been shown (François et al., 2002). François et al. (2003) also demonstrated that tert-butyl formate (TBF), the intermediate between MTBE and TBA, exerted inhibitory effects on MTBE and TBA oxidation.
Inefficient MTBE biodegradation could also be the result of a loss of energy (Fortin et al., 2001), probably related to a lack of reduced co-enzymes and protons, thus de-coupling the proton pump and leading to a shortage of ATP. Also, the MBTE pathway has an unusually high requirement in reducing equivalents (Salanitro, 1995). The characterization of the enzymes involved in the MTBE degradation pathway is crucial to understand the limitations to MTBE degradation, but it has not yet been documented. To the best of our knowledge, we are the first to describe the isolation and characterization of a gene cluster, designated the mpd cluster, containing genes encoding an alcohol dehydrogenase and an aldehyde dehydrogenase, which are both involved in the MTBE metabolic pathway of M. austroafricanum IFP 2012. Functional heterologous expression was carried out in Mycobacterium smegmatis mc2 155 and the involvement of these enzymes in the specific degradation of 2-M1,2-PD to HIBA is demonstrated.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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. M. austroafricanum IFP 2012 was grown at 30 °C in Luria–Bertani (LB) medium, or in the mineral medium (MM) previously described (Piveteau et al., 2001), supplemented with MTBE, TBA, 2-M1,2-PD, HIBA or glucose as the sole carbon source. The headspace volume was sufficient to prevent any oxygen limitation during growth. Growth was followed by measuring the OD600. The substrate or metabolite concentrations were measured using filtered samples by GC or HPLC. M. smegmatis mc2 155, Escherichia coli DH10B and respective recombinant clones were grown in LB supplemented with kanamycin (20 µg ml–1) or ampicillin (100 µg ml–1), as required.
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), except for the migration conditions (150 V or 7·5 V cm–1) during the first hour, and a constant current of 33 mA for the last 5 h.
Peptide sequencing of proteins induced during growth in the presence of MTBE.
For internal sequencing, MTBE-induced protein bands from the MTBE profile on SDS-PAGE were cut out. An ‘in-gel’ reduction (DTT) and an alkylation by iodoacetamide of the disulfide groups (Jenö et al., 1995) were performed prior to modified sequencing grade trypsin digestion (Hellman et al., 1995) (Promega). Edman degradation was performed according to Hewick et al. (1981). The N-terminal sequences of the isolated proteins, previously transferred onto a Problott membrane (Applied Biosystems), were obtained by automated Edman degradation, using the sequencing protocol described above. For internal peptide sequencing, about 1 pmol of the peptide was loaded onto a model 494 cLC Procise sequencer, employing the general protocol (Hewick et al., 1981), and a standard program using liquid-phase trifluoroacetic acid was employed for further sequencing.
Design of degenerate oligonucleotide primers.
Degenerate primers were designed from N-terminal or internal peptide sequences of specific MTBE-induced proteins. The deduced DNA sequence was obtained according to universal codon usage and primers were created in the least degenerate part of each DNA sequence. Correlations between genome composition (in terms of G+C content) and usage of particular codons and amino acids have been widely reported and Giegel et al. (1990) demonstrated that the presence of an adenine at the third position in codons is rare in coding sequences of mycobacteria. When adenine was the base at this position, it was randomly replaced with one of the other three nucleotides to reduce primer degeneracy. Annealing temperatures used for PCR experiments were calculated at 2 °C lower than the lowest melting temperature found in the primer pool.
Total DNA extraction.
Total DNA was isolated from M. austroafricanum IFP 2012 according to Pospiech & Neumann (1995), with the following modifications: a 50 ml culture grown in LB medium to an OD600 of 0·7 was centrifuged (8000 g, 4 °C, 15 min) and resuspended in 5 ml SET buffer (75 mM NaCl, 25 mM EDTA, 20 mM Tris/HCl, pH 8·0). Lysozyme (final concentration 1 mg ml–1) prepared in 10 mM Tris/HCl, pH 8·0, and 500 µl lysostaphin at 50 µg ml–1 (Sigma), prepared in 10 mM Tris/HCl, pH 7·5, were added to the cell suspension before incubation at 37 °C for 1 h. After SDS/proteinase K treatment, an extraction step with NaCl/cetyltrimethylammonium bromide (CTAB) replaced the addition of 0·33 vols 5 M NaCl; 0·25 vols 5 M NaCl (1 M final concn) and 0·13 vols 10 % CTAB/0·7 M NaCl (pre-heated to 65 °C) were added sequentially and the mixture was incubated for 10 min at 65 °C. Then, 1 vol. CHCl3 was added and the subsequent steps were carried out according to the original protocol.
Cloning and sequence analysis of the M. austroafricanum IFP 2012 mpd gene cluster.
The plasmids used for the cloning are described in Table 1
and the primers pairs used at the different steps are listed in Table 2. Southern blot analysis was carried out on total DNA of M. austroafricanum IFP 2012 restricted with different enzymes and probed with a 204 bp PCR fragment (mpd probe) labelled with digoxigenin-dUTP (DIG DNA Labelling and Detection Kit; Roche Diagnostics). The size of the SmaI-positive band was estimated to be between 5 and 6 kbp; this region of the gel was excised, the DNA was purified and then used to construct a DNA library in pBluescript II KS+ (Stratagene) cut with SmaI. For transformation, E. coli DH10B cells were treated chemically according to Hanahan et al. (1991) and transformants were screened by colony hybridization using the mpdB probe. Plasmid (pKS1) from one probe-positive transformant was extracted by a mini-prep procedure (QIAprep Spin Miniprep kit; Qiagen). The presence of the SmaI insert was checked with a new digestion with SmaI and this revealed a unique band corresponding to the size of the linearized plasmid. This could be explained by the presence of a damaged DNA insert containing only one SmaI site.
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To obtain the upstream mpdC gene sequence, a 904 bp PCR fragment, spanning part of the mpdC gene obtained by PCR using primers MF1 and MR1, was used as a probe (mpdC probe) for Southern hybridization to identify the 5574 bp PstI fragment. The 5–6 kbp PstI fragments were cloned in pBluescript II KS+, as described above, to obtain plasmid pKS3. The entire inserts of the resulting positive recombinant plasmids were sequenced on both strands by primer walking with a T7 DNA-sequencing kit (Applied Biosystems), using the ABI Prism 377 automated fluorescence sequencer (Applied Biosystems). To minimize artifacts that are often associated with high G+C content sequences, the following modifications were incorporated. Prior to amplification and the addition of the Big Dye Terminator, the DNA was denatured at 98 °C for 5 min. A mixture of Big Dye Terminator v3.1 and dGTP was also used in a ratio of 3 : 1. The nucleotide sequences, the ORFs determined by the ORF finder software (http://www.ncbi.nlm.nih.gov/gorf/gorf.html), and the corresponding amino acid sequences were compared with the EMBL, SWISS-PROT and GenBank databases using BLASTN and BLASTX from the National Center for Biotechnology Information (Altschul et al., 1997).
Construction of an mpd gene expression vector and transformation in M. smegmatis mc2 155.
The mpd gene cluster was introduced into pCL4D (Picardeau et al., 2000) using a two-step procedure. In the first step, the 4411 bp NotI fragment of pKS2 (Table 1) was blunt-ended with T4 DNA polymerase (NEB) and cloned into the HincII site of pCL4D. Transformation was carried out in E. coli DH10B and the transformants were selected on LB agar containing kanamycin (20 µg ml–1). The screening was performed by PCR amplification with primers F-M2 and R-M2 (Table 2, Fig. 2) to identify the transformants harbouring the positive recombinant plasmid p4D1. In the second step, the 5574 bp PstI fragment of pKS3, carrying orf3, mpdR and part of the mpdC gene, was introduced into p4D1 cut with PstI, from which a 950 bp fragment was removed to finally generate p4D2 (Fig. 2). The orientation of the PstI fragment in the recombinant plasmid was also screened by PCR using primers F-M3 and R-M3 (Table 2, Fig. 2) to isolate the final p4D2 recombinant plasmid carrying the entire mpd gene cluster. The mpd gene cluster inserted into the p4D2 plasmid was introduced by electroporation into M. smegmatis mc2 155 as follows. Cells of M. smegmatis mc2 155 were prepared for electroporation according to Parish & Stoker (1998), with some modifications: M. smegmatis mc2 155 was grown at 37 °C to exponential phase (OD600 
0·7) in LB medium supplemented with 0·1 % Tween 80 to reduce formation of aggregates. Pellets were washed five times, sequentially, with 1, 0·5, 0·05 and 0·01 vols washing buffer (10 %, v/v, glycerol/0·1 % Tween 80), resuspended in 4 ml 10 % glycerol solution and 80 µl aliquots were stored at –80 °C. The competent cells (80 µl) were mixed with 1 µg pCL4D or p4D2 in a 0·2 cm gap cuvette. The electroporation conditions were 2·5 kV, 25 µF and 200 
in a Gene Pulser (Bio-Rad Laboratories). Electroporated cells were recovered in 0·4 ml SOC liquid medium (Ausubel et al., 1990) and incubated for 2 h at 37 °C before plating. A transformant, M. smegmatis mc2 155(p4D2), was isolated on LB medium containing kanamycin (20 µg ml–1). M. smegmatis mc2 155 was also electroporated with pCL4D without any insert and the resulting transformant M. smegmatis mc2 155(pCL4D) was used in control experiments.
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RNA extraction.
All glassware and solutions were made RNase-free according to standard procedures (Sambrook & Russell, 2001). Total RNA of M. austroafricanum IFP 2012 was extracted from a 20 ml culture grown to an OD600 0·7 in MM medium supplemented with the required substrate. After incubation on ice for 30 min, cells were harvested (8000 g, 10 min, 4 °C) and kept on ice. Lysozyme (Roche Diagnostics) (600 µl of 3 mg ml–1), prepared in 10 mM Tris/HCl, pH 8·0, and lysostaphin (Sigma) (600 µl of 50 µg ml–1), prepared in 10 mM Tris/HCl, pH 7·5, were added to the cell pellet and incubated at 37 °C for 10 min. RNAwiz (10 ml) (Ambion) was added and the mixture was vigorously mixed for 15 s in a vortex mixer equipped with a vortex adapter (Ambion). The resulting mixture was divided into 2 ml tubes (1400 µl per tube) containing 250 mg zirconium-silica beads (0·1 mm diameter) and mixed for 10 min on the vortex mixer. After centrifugation (13 000 g, 4 °C, 5 min), the bacterial lysates were recovered from each tube, transferred to a clean 2 ml tube and 0·2 vols CHCl3 was added. The tubes were then vortexed for 30 s, incubated at room temperature (10 min), centrifuged (13 000 g, 4 °C, 5 min) and the aqueous supernatants were transferred to another 2 ml tube. RNA extracts were precipitated by sequentially adding 0·5 vols H2O treated with diethyl pyrocarbonate (DEPC), 1 vol. 2-propanol and 0·02 vols RNase-free glycogen at 5 mg ml–1 (Ambion), mixed well and incubated at room temperature for 10 min. The RNA was pelleted (13 000 g, 4 °C, 15 min), washed twice with 70 % ethanol, dried for 30 s (DNA Speed Vac; Savant) and resuspended in sterile tubes in 25 µl DEPC-treated H2O. The quality of the pooled RNA was assessed by agarose gel electrophoresis (1 % agarose gel in 0·5x TBE). RNA was stored at –80 °C, precipitated with 0·1 vols 3 M sodium acetate, pH 7·0, and 2·5 vols 95 % ethanol.
DNase treatment and RT-PCR.
A 2 µg quantity of RNA extracted from M. austroafricanum IFP 2012 was treated with DNA-free (Ambion), according to the manufacturer's instructions. DNA removal was assessed by PCR using primers specific for the M. austroafricanum IFP 2012 mpd gene cluster (Table 2). The final concentration of RNA was quantified at 260 nm using NanoDrop (NanoDrop Technologies). RT-PCR was performed using the Qiagen OneStep RT-PCR kit, with 120 ng RNA. A positive PCR control reaction, i.e. with 100 ng M. austroafricanum IFP 2012 total DNA, and a negative RT-PCR control for each set of primers used on every RNA preparation, were also prepared. The three RT-PCR primer pairs tested were: RT-PCR FI/RI, RT-PCR FII/RII and RT-PCR FIII/RIII (Table 2).
RT-PCR conditions were as follows. Reverse transcription step: 50 °C, 30 min. Positive, negative and (–) RT-PCR controls were then added, at 50 °C, before PCR amplification cycles. PCR amplification step: 95 °C, 15 min; 35 cycles of 94 °C, 30 s; 55 °C, 1 min; 72 °C, 1 min, followed by a final extension at 72 °C for 10 min. Samples were then stored at 4 °C until they were analysed.
Analytical procedures.
MTBE, TBA and 2-M1,2-PD were quantified by flame-ionization detection on a Varian 3300 gas chromatograph (Varian) equipped with a 0·32 mmx25 m Porabond-Q capillary column (J&W Scientific), using a gradient ranging from 105 to 210 °C at 10 °C min–1, followed by a steady temperature of 210 °C for 20 min. Helium (1·6 ml min–1) was used as the carrier gas. Samples, filtered through 0·22 µm filters (Prolabo), were injected without further treatment. HIBA was quantified in filtered samples by HPLC (Metrohm), using an anion column (Dual 2, 75 mmx4·6 mm) and a conductivity detector (Metrohm 732). The eluent, a mixture of 1·8 mM Na2CO3 and 1·7 mM NaHCO3, was previously flushed by a Spectra-Physics SCM 400 vacuum flusher and the flow rate was 1 ml min–1.
Chemicals.
MTBE, TBA and HIBA were of the highest purity available (Sigma-Aldrich) and 2-M1,2-PD was a kind gift from Dr Sergio Revah (Universidad Iztapalpa, Mexico City, Mexico).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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(a) and (b). Five overexpressed proteins, with molecular masses of 67, 64, 62, 55 and 50 kDa, were detected on the 10 % polyacrylamide gel by comparing the protein patterns after growth in the presence of MTBE and glucose. Five other induced proteins, with molecular masses of 27, 23, 17, 14 and 11, were detected on the 15 % polyacrylamide gel by comparing the protein patterns after growth in media containing MTBE and glucose.
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N-terminal and internal protein sequencing was carried out on induced proteins recovered from MTBE profiles, as described in Methods. Peptide sequences obtained were analysed using ‘BLAST search for short, nearly exact matches' (Altschul et al., 1997). The N-terminal sequences of the 62 and 11 kDa bands KIIAYDEEAR and ASVNIKPLEDKILV were 95 % similar to the molecular chaperones of mycobacteria GroEL and GroES, respectively. A band, KNPLFLDEQLTRAEFQ, corresponding to an internal sequence of the 67 kDa protein, was homologous to the DnaK molecular chaperone of Mycobacterium tuberculosis. Two internal sequences of 16 and 30 aa, KQRGWAYDPNVRGLPE and STEHGLEGTIDWPISYEELAPYYDENDAIY (François et al., 2002), belonging to the 64 kDa protein, were obtained. The GenBank homology search revealed that the 30 aa sequence had a significant similarity (93 %) with several enzymes of the (glucose–methanol–choline) GMC oxidoreductase family (Cavener, 1992), which includes oxidases and dehydrogenases.
The N-terminal sequences of the 23 kDa protein (SRTDILAATQAAGAPIEQVV) and the 17 kDa protein (NXVAVQQDTREAVGE) had similarities with a ribulose-1,5-bisphosphate carboxylase (or Rubisco) of Chloromonas spp. (63 % similarity) and the pyrene dioxygenase small subunit of Mycobacterium sp. PYR-1 (80 % similarity), respectively. An internal amino acid sequence from the 50 kDa induced protein (KSFHPFPRAMMGPGAHEMVGPEALK; François et al., 2002) matched well (61 % similarity) with the partial sequence of a protein from Mycobacterium gastri, known as methanol : N,N'-dimethylnitrosoaniline (NDMA) oxidoreductase (Bystrykh et al., 1993).
Peptide sequences obtained from the other MTBE-induced proteins did not show any significant similarity with known enzymes.
Amplification of a DNA sequence of the 64 kDa protein (mpdB gene)
The 16 and 30 aa sequences obtained from the 64 kDa protein were used to design the degenerate oligonucleotide primers MadF1 and MadR1 (Table 2). A 204 bp PCR product, designated the mpd probe, was obtained and sequenced. The peptide sequence correlated with the deduced amino acid sequence of the PCR product, which revealed a partial ORF corresponding to 83 aa. BLAST analysis of this new peptide sequence confirmed the similarity of the 64 kDa protein with flavoproteins of the GMC oxidoreductase family, which includes a variety of enzymes, such as choline dehydrogenase (CDH) and methanol oxidase (MOX). In the case of the 64 kDa protein, the best alignment was obtained (61 % similarity) with a putative choline dehydrogenase from Bradyrhizobium japonicum USDA110. The amino acid sequence of this choline dehydrogenase was investigated to detect conserved domains that could be used to obtain a larger DNA sequence. We were able to detect the Walker-type-A motif GXGXXG, which was first detected by Cavener (1992), described by Wierenga et al. (1985) and now known to be an ATP/GTP binding motif. This motif was used to design a new forward degenerate primer MadF2 (Table 2). PCR amplification yielded an unexpected 604 bp DNA fragment, resulting from the new primer annealing upstream of the expected annealing position. The PCR fragment, designated the mpdB probe, was subsequently used for colony hybridizations.
Cloning of the M. austroafricanum IFP 2012 genomic DNA (gDNA) fragment containing the mpd gene cluster
Southern blot analysis of SmaI-digested total DNA of M. austroafricanum IFP 2012, using the 204 bp mpdB probe, revealed a 6 kbp probe-positive fragment (data not shown). To clone the mpdB-positive fragment, a DNA library was constructed in pBluescript II KS+ and screened by colony hybridization using the mpdB probe. The plasmid pKS1 was isolated from a positive transformant. All plasmid preparations of purified clones showed several bands on agarose gels. For each gel, we observed a major band of 3 kbp, corresponding to the empty linearized plasmid, and minor bands below the expected size, suggesting that the insert introduced into pBluescript II KS+ was highly unstable. The sequencing was carried out after gel extraction of the SmaI-linearized pKS1. Preliminary sequence analysis of the insert in pKS1 revealed that this 5676 bp gDNA fragment did not contain the 5' upstream part of the mpdC gene. Further Southern blot and colony hybridization analyses were then performed to obtain a 5574 bp PstI fragment, using a 904 bp mpdC probe. This probe was obtained using the MF1/MR1 primer pair designed in the mpdC gene fragment included in the SmaI insert (see Fig. 2 for localization). The pKS3 plasmid containing the 5574 bp gDNA fragment was isolated from a transformant showing positive hybridization.
Analysis of the 10 327 bp PstI–SmaI gDNA fragment
Compilation of the SmaI and PstI inserts, both containing the mpdC probe, resulted in a 10 327 bp gDNA sequence of M. austroafricanum IFP 2012. The DNA and deduced amino acid sequences of this overlapping PstI–SmaI fragment revealed seven ORFs. We identified six complete genes: orf3, mpdC (encoding a hydroxyisobutyraldehyde dehydrogenase), orf1 (encoding an MTBE-induced polypeptide), mpdB (encoding a 2-M1,2-PD dehydrogenase), orf2 (encoding a membrane protein) and mpdR (encoding a putative regulatory protein). The genes were all in the same orientation, except mpdR, which was located between orf3 (one transmembrane domain protein) and mpdC (Fig. 2). BLAST comparisons of each putative coding sequence with known proteins are presented in Table 3.
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54-dependent transcriptional regulator, but only in the N-terminal moiety of the protein.
The mpdC gene, encoding a 55 kDa protein, had a similar size and the highest amino acid sequence identity (45 % identity/61 % similarity) with an NAD-dependent aldehyde dehydrogenase of Burkholderia fungorum. The CD-search analysis revealed a strong alignment with the ‘pfam00171.11 aldedh’ domain (E value=1e–123), which has been described in the Pfam database of protein families (Bateman et al., 2002), and corresponds to an NADP-dependent aldehyde dehydrogenase family. To predict the secondary structure of the MpdC protein sequence, we used the two prediction servers PSIPRED (McGuffin et al., 2000) and SSPro (Pollastri et al., 2002). Both of these servers are known for their comparatively good performance of over 75 % accuracy on average (Rost & Eyrich, 2001), although they are based on different computational methods. Comparison of MpdC with the well-described structure of the betaine aldehyde dehydrogenase (BetB) of cod liver (Johansson et al., 1998) revealed the presence of 27 of the 33 secondary structures previously determined in BetB of cod liver (Fig. 3a).
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The mpdB gene, located immediately downstream of orf1, encoded a 64 kDa protein. Primary DNA sequence analysis correlated with the N-terminal peptide sequence obtained (MTTSADQTDVLVIG; this study) and confirmed the presence of a GTG start codon. This start codon is not frequently found, but has been described in coding sequences of mycobacteria (Hu & Coates, 1999; Kamalakannan et al., 2002; Misra et al., 1996; Revel-Viravau et al., 1996). The amino acid sequence, which includes those of the peptide fragments directly sequenced from the 64 kDa protein induced in the presence of MTBE, had the best alignment (42 % identity/56 % similarity) with a putative GMC oxidoreductase of B. japonicum. The CD-search analysis revealed a strong alignment with a cluster of orthologous domains (COG; Tatusov et al., 1997): the ‘COG23031 BetA’ (E value=4e–49). The amino acid consensus sequence of COG2303 characterizes choline dehydrogenases and related flavoproteins, which are involved in the metabolism of choline. The alignment result of the most conserved region of MpdB with BetA of E. coli (Lamark et al., 1991), Sinorhizobium meliloti (Pocard et al., 1997) and COG2303 is shown in Fig. 3(b). We were able to detect the four highly conserved representative regions of the FAD-binding domain of choline dehydrogenases (Kiess et al., 1998). This FAD-binding domain contains the Walker-type-A motif previously detected and a characteristic 
-sheet domain which was revealed in MpdB by the secondary structure prediction.
The last ORF of the mpd cluster, orf2, encoded a 25 kDa protein, most closely related to the ‘COG3104’ domain (37 % identity/48 % similarity), including a putative di/tri-peptide permease of R. gelatinosus PM1. Hydropathy analysis of the protein revealed the presence of five transmembrane peptide segments, suggesting that it is located in the membrane. This protein was not detected by SDS-PAGE of the cytoplasmic extract of M. austroafricanum IFP 2012, which did not include any membrane polypeptides.
Nucleotide sequence analysis of the mpd gene cluster showed three short intercistronic regions of 68 (mpdC–orf1), 25 (orf1–mpdB) and 89 (mpdB–orf2) nt, including, in each case, a potential Shine–Dalgarno sequence.
An incomplete ORF, the tnpA gene, was found at the 3' end of the cloned gDNA fragment (Fig. 2). The ORF encodes an amino acid sequence which strongly matched (47 % identity/59 % similarity) with two putative transposases of Rhodococcus erythropolis. Analysis of the orf2–tnpA intergenic region showed that it was divided into two parts: the upstream sequence (110 nt) had an abnormally high A+T content (50 %), while the 3' region showed the high G+C content (66 %) usually found in mycobacteria.
Finally, orf3, which was upstream of mpdR (Fig. 2), did not show significant homology with any known protein in either the protein or the DNA database, but hydropathy analysis revealed a single C-terminal transmembrane domain.
Heterologous expression of the mpd gene cluster in M. smegmatis mc2 155
The isolated 9060 bp fragment of M. austroafricanum IFP 2012 gDNA, deleted for the tnpA incomplete gene, was cloned in the pCL4D shuttle vector to yield the p4D2 expression plasmid, which was used to transform M. smegmatis mc2 155. The empty plasmid pCL4D was also used to transform M. smegmatis mc2 155. Degradation capacities of the resulting recombinant strains M. smegmatis mc2 155(p4D2) and M. smegmatis mc2 155(pCL4D) were determined using resting cells after growth on LB medium containing kanamycin. No degradation of MTBE, TBA and HIBA was observed with either strain. M. smegmatis mc2 155(p4D2) degraded 2-M1,2-PD and a stoichiometric production of HIBA was observed (Fig. 4). The maximum rate of 2-M1,2-PD degradation by strain M. smegmatis mc2 155(p4D2) was 2·34±0·41 µmol min–1 (g dry weight)–1 and the maximum rate of HIBA production was similar at 1·89±0·26 µmol min–1 (g dry weight)–1. This result clearly showed that the function of the cloned cluster was the oxidation of 2-M1,2-PD to HIBA by two enzymic steps, which were catalysed by 2-M1,2-PD dehydrogenase (MpdB) and hydroxyisobutyraldehyde dehydrogenase (MpdC).
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). No degradation was observed either in resting cells of M. smegmatis mc2 155(pCL4D), containing the vector without insert, or in the abiotic control, demonstrating the specific role of the mpd gene cluster in the conversion of 2-M1,2-PD to HIBA via hydroxyisobutyraldehyde (Fig. 5). The detection of hydroxyisobutyraldehyde by GC/MS was not successful, probably because the rate of 2-M1,2-PD degradation was equal to the rate of HIBA production, preventing accumulation of this intermediate.
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, respectively). We used three different primer pairs designed to amplify (i) an internal sequence of mpdR (RT-FI/RT-RI primer pair), (ii) the overlapping sequence mpdC–orf1 (RT-FII/RT-RII primer pair), and (iii) the overlapping sequence mpdB–orf2 (RT-FIII/RT-RIII primer pair). These primer pairs are described in Table 2. Strong RT-PCR amplification with both mpdC-orf1 and mpdB-orf2 primer pairs was observed using RNA extracted from M. austroafricanum IFP 2012 cells grown on TBA. In addition, we also obtained amplification using the mpdR primers (Fig. 6a). These results clearly showed that transcription of the mpd gene cluster was induced after growth on TBA.
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). This could suggest a basal transcription level of the mpd gene cluster under non-selective conditions. |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONREFERENCES |
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We characterized proteins specifically induced after growth in the presence of MTBE (Fig. 1) and determined the internal or N-terminal amino acid sequences of these proteins. The DNA sequences deduced from these peptide sequences were used to design degenerate oligonucleotide primers to obtain PCR amplification products. The products were labelled and used as probes in Southern analysis on digested total DNA of M. austroafricanum IFP 2012 to isolate and clone a 10 327 bp gDNA fragment containing seven ORFs (six complete and one incomplete, Fig. 2). A 9060 bp fragment was then cloned in pCL4D and transferred into M. smegmatis mc2 155. The functional heterologous expression of these genes isolated from M. austroafricanum IFP 2012 in M. smegmatis mc2 155 was also shown for the first time, to the best of our knowledge, by determining that the products of the mpd genes, including an alcohol dehydrogenase (MpdB) and an aldehyde dehydrogenase (MpdC), catalysed the conversion of 2-M1,2-PD to HIBA (Fig. 4). This activity was inducible by 2-M1,2-PD, as it occurred after a 4 h lag phase and did not occur when chloramphenicol, which inhibits translation, was present. The 2-M1,2-PD was previously identified as the product of TBA oxidation in M. austroafricanum IFP 2012 (Lopes Ferreira et al., 2005) and our latest results show that the mpd genes are involved in MTBE catabolism in M. austroafricanum IFP 2012. In addition, we also detected the correlated overexpression of 64, 55 and 27 kDa proteins in M. austroafricanum IFP 2012 grown in the presence of MTBE, as shown in Fig. 1(a) and (b), and these proteins corresponded to the products of the mpdB, mpdC and orf1 genes, respectively.
Sequence alignment of MpdB and MpdC with protein databases showed high similarities with BetA and BetB, respectively, which are two enzymes responsible for the synthesis of glycine betaine from choline in several bacteria (Csonka & Epstein, 1996; Lamark et al., 1991; Landfald & Strom, 1986; Pocard et al., 1997). The MpdB protein showed the highest similarity with a choline dehydrogenase of B. japonicum, which is a strain unable to degrade choline (Boncompagni et al., 1999), and this was also the case in M. austroafricanum IFP 2012 (data not shown).
Structural alignment of MpdB with choline dehydrogenases is described in Fig. 3(b). This alignment showed the conservation of several consensus patterns required in the FAD-binding domain, such as a –
–-fold domain, a flavin-attachment loop, -sheet domains and amino acids involved in the active sites. Structural alignment of MpdC with betaine aldehyde dehydrogenases (Fig. 3a) showed a strong conservation of both the amino acid sequences and the secondary structures of all the related proteins that were examined. Among the putative proteins encoded by the mpd genes, only MpdB and MpdC were closely related to functional alcohol dehydrogenase and aldehyde dehydrogenase enzymes, respectively.
Since the conversion of 2-M1,2-PD to HIBA requires the sequential action of an alcohol dehydrogenase and an aldehyde dehydrogenase, we can infer from these results that these two enzymes are specifically involved in this conversion in the MTBE metabolic pathway (Fig. 5).
Two distinct phases in the MTBE pathway can be identified during the degradation from MTBE to HIBA. The first one, from MTBE to 2-M1,2-PD, includes two oxidation steps catalysed by monooxygenases: (i) MTBE to tert-butoxymethanol, and (ii) TBA to 2-M1,2-PD. This part of MTBE catabolism can be written as the sum of five enzymic reactions, including the mineralization of formate generated as a co-product by the hydrolysis of TBF to TBA: C5H12O+2O2
C4H10O2+H2O+CO2.
This upper part of the MTBE degradation pathway consumes reduced coenzyme equivalents. Even though NADH is regenerated by formate dehydrogenase during the conversion of formate to carbon dioxide, no ATP can be produced, resulting in a loss of energy. That is one of the reasons why MTBE-degrading micro-organisms have to be methylotrophic, which is the case for M. austroafricanum IFP 2012 (François et al., 2002). In the lower part of the pathway, the oxidation of 2-M1,2-PD to HIBA by the action of two successive dehydrogenases [C4H10O2+H2O+2NAD(P)+C4H8O3+2NAD(P)H+2H+] produces reduced coenzyme equivalents that can be used in the above oxidation reactions. The requirement for reducing equivalents could explain (i) the strong expression of the enzymes encoded by the mpd cluster observed in the MTBE protein profile (Fig. 1), and (ii) the decrease in the MTBE degradation rate observed during growth of M. austroafricanum IFP 2012 (François et al., 2002). The efficiency of the degradation of 2-M1,2-PD to HIBA explains why hydroxyisobutyraldehyde was not be detected as a degradation intermediate, since HIBA was produced at the same rate as 2-M1,2-PD was degraded [1·89±0·26 µmol min–1 (g dry weight)–1 and 2·34±0·41 µmol min–1 (g dry weight)–1, respectively]. Moreover, the production of reduced co-enzyme equivalents is linked to the production of ATP and it has been proposed that the energy loss during growth in the presence of MTBE could be a key limiting factor in its catabolism (Fayolle et al., 2003; Fortin et al., 2001, Salanitro, 1995). It is known that the imbalance in reduced co-factors generated during the oxidation steps of xenobiotic biodegradation can limit growth of micro-organisms on these compounds (VanBriesen, 2001).
A high level of mRNA transcription was observed by RT-PCR following growth on TBA, clearly showing the specific induction of the mpd genes. This correlated with the results obtained from SDS-PAGE analysis.
The mpd genes appeared to be regulated by mpdR, which encodes a putative 54-dependent transcriptional regulator (Burgess et al., 1969). However, alignments were found only in the N-terminal part of the corresponding protein. Moreover, two motifs that are usually conserved, i.e. the N-terminal ATP-binding motif (Austin et al., 1991; Morett & Segovia, 1993), and the C-terminal helix–turn–helix DNA-binding domain (Austin & Dixon, 1992), were not detected. Nevertheless, it is noteworthy that genes encoding 54 were not found in the genomes of several recently sequenced Gram-positive strains that had a high-G+C content (Buck et al., 2000). DNA analysis of the mpd cluster also revealed short intergenic regions, including Shine–Dalgarno sequences (Draper, 1996). A GUG start codon was present in the coding sequence of the mpdB gene and this is unusual (Ma et al., 2002). Due to the short intergenic sequences separating the regulatory mpdR gene from the other mpd genes, promoters are probably overlapping, which should contribute to the complexity of the regulation. Røkenes et al. (1996) have shown that the betIBA operon of E. coli contains similar overlapping promoters.
Of interest was the specific expression of the molecular chaperones DnaK and GroEL/ES which were induced during growth in the presence of MTBE. Molecular chaperones are involved in a myriad of functions, including nascent protein folding and assembly, refolding of protein quaternary structure (Lopes Ferreira & Alix, 2002), synthesis of mycolic acids, translocation across membranes, or modulation of heat-shock responses, which can be considered as housekeeping functions, and a defence strategy against stress (for review see Alix, 2004). The combination of the high folding demand of newly synthesized proteins and the above-mentioned imbalance in reduced co-factors during growth in the presence of MTBE could explain the stimulation of these molecular chaperones in M. austroafricanum IFP 2012.
From a global perspective, our results showed that growth in the presence of MTBE induces ten cytoplasmic proteins. Seven of these proteins were characterized: (i) proteins of 64, 55 and 27 kDa encoded by the mpdB, mpdC and orf1 genes, respectively, described in this study; (ii) proteins of 67, 63 and 11 kDa corresponding to DnaK and GroEL/ES molecular chaperones, respectively; and (iii) a protein of 50 kDa, which is the methanol : NDMA oxidoreductase detected in Mycobacterium gastri (Bystrykh et al., 1993) and several other mycobacteria (Park et al., 2003). This latter protein was detected in crude extracts of M. austroafricanum IFP 2012 after growth on HIBA and 2-propanol (François, 2002), suggesting its involvement in assimilation of 2-propanol, a putative intermediate of the MTBE metabolic pathway. The remaining induced proteins corresponded to: (i) a 24 kDa protein, which matched with the Rubisco involved in carbon dioxide fixation; (ii) a 17 kDa protein, which was closely related to the naphthalene dioxygenase small subunit from Mycobacterium sp. PYR-1 (Khan et al., 2001); and (iii) an uncharacterized 14 kDa protein. The role of the latter three proteins in the MTBE catabolic pathway has not yet been determined.
At present, the nature of the monooxygenases catalysing the oxidation of MTBE and TBA has not been elucidated, as none of the strongly induced soluble proteins have shown similarities with known monooxygenases. The hypothesis that the monooxygenase responsible for MTBE and TBA oxidation might be located in the membrane deserves further investigation, since we did not analyse this cell compartment. In this regard, the membrane-bound alkane hydroxylase could be a candidate, since several bacterial strains are able to degrade MTBE by cometabolism after growth on alkanes (Garnier et al., 1999; Smith & Hyman, 2004; Smith et al., 2003).
The products of the mpd genes involved in the biodegradation of 2-M1,2-PD play a key role in the growth of M. austroafricanum IFP 2012 in the presence of MTBE. The mpdB and mpdC sequences could provide new molecular tools to undertake studies on the distribution of MTBE-degrading bacteria. These tools should help in estimating the capacity for natural attenuation of MTBE-contaminated ecosystems.
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Received 12 October 2005;
revised 11 January 2006;
accepted 16 January 2006.
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