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Previous Article | Next Article 
Applied and Environmental Microbiology, April 2007, p. 2580-2591, Vol. 73, No. 8
0099-2240/07/$08.00+0 doi:10.1128/AEM.02074-06
Copyright © 2007, American Society for Microbiology. All Rights Reserved.
Department of Biological Sciences, University of Warwick, Coventry CV4 7AL, England, United Kingdom
Received 2 September 2006/ Accepted 12 February 2007
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INTRODUCTION |
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Growth on DMS as a carbon source has been described for a range of prokaryotes, including anaerobic degradation by methanogens (28) and sulfate-reducing bacteria (48). Aerobic bacterial DMS oxidation was first demonstrated for some members of the genera Hyphomicrobium and Thiobacillus (9, 24, 40a, 47). In these bacteria, DMS monooxygenase was identified as a key enzyme in DMS metabolism, producing methanethiol and formaldehyde. DMS monooxygenase activity was also found in Hyphomicrobium sp. strain S growing on DMSO (9) and in strains of Hyphomicrobium sulfonivorans that were isolated on dimethyl sulfone as the carbon source (4, 37).
The phylogenetic diversity of marine DMS-degrading prokaryotes is still largely unexplored. Alphaproteobacteria, especially members of the Roseobacter clade, have often been implicated in the metabolism of organosulfur compounds in the marine environment (16, 38, 56), but it is not clear whether these bacteria are able to grow on DMS. Marine isolates growing on DMS as the carbon source, obtained from marine sediments, included Rhodovulum sulfidophilum SH1, Thiobacillus sp. strain ASN-1, Thiobacillus thioparus T5, Thiocapsa roseopersicina M11, Methylophaga sulfidovorans, and the unidentified isolate BIS-6 (17, 23, 36, 50, 51). Less is known about the diversity of DMS-degrading bacteria in the pelagic marine environment. Recently, Vila-Costa and colleagues (49) reported the detection of Methylophaga spp. by denaturing gradient gel electrophoresis (DGGE) and clone library analysis of DMS enrichment cultures from seawater samples. Unfortunately, isolates were not obtained and so the assumption that the detected populations of Methylophaga were indeed able to grow on DMS could not be substantiated. Previously reported DMS-degrading bacterial isolates from pelagic marine samples that could grow on DMS were not identified by sequencing of 16S rRNA genes (18, 20), further highlighting the need to cultivate and identify DMS-degrading bacteria from seawater.
Given the phylogenetic diversity of DMS-degrading bacteria thus far identified, and the fact that closely related isolates of DMS-degrading strains may be unable to grow on DMS, the identification of DMS-degrading populations in environmental samples based on 16S rRNA genes is difficult. Functional molecular markers, i.e., PCR primers and probes targeting genes encoding key enzymes of DMS degradation pathways, would therefore be invaluable tools with which to study the abundance and distribution of DMS-degrading bacteria in environmental samples and to characterize the diversity of genes and enzymes involved in this globally relevant process. However, the genes encoding DMS monooxygenases, DMS methyltransferases, or other key enzymes of DMS metabolism from organisms growing on DMS as a carbon source have not yet been identified.
The aims of this study were (i) to identify bacterial populations in marine DMS-degrading enrichment cultures, (ii) to identify isolates capable of growth on DMS, and (iii) to identify polypeptides involved in metabolism of DMS. These were achieved by analyzing enrichment cultures by denaturing gradient gel electrophoresis analysis, sequencing 16S rRNA genes of isolates, testing the ability of isolates to grow on DMS, and characterizing the genetic diversity of DMS-degrading Methylophaga isolates by BOX-PCR (42). Finally sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) analysis of cell extracts from biomass of a Methylophaga isolate revealed polypeptides induced during growth on DMS which were identified by mass spectrometry techniques and N-terminal sequencing.
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MATERIALS AND METHODS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Enrichment cultures were also set up using unialgal Emiliania huxleyi cultures as the inoculum. Microscopic observation showed that none of the E. huxleyi strains was axenic (M. Cox, personal communication). Two-milliliter culture aliquots of E. huxleyi strains 92A, 371, 373, 373UEA, and 1516 were pooled and gently vacuum filtered through a 0.2-µm-pore SUPOR membrane filter (Pall, Farlington, United Kingdom). The filter was rinsed by filtering 15 ml of MAMS through the membrane before the biomass retained on the filter was resuspended in 4 ml of MAMS, and 400 µl of the suspension was used to inoculate 25 ml of MAMS in sealed, crimp-top vials. A second culture of strain 1516 that had previously been axenic was used separately as an inoculum due to its markedly higher turbidity.
Aliquots of 2.5 ml from each of six different methyl halide-degrading enrichment cultures (44) were pooled and used to inoculate enrichment cultures which were amended with DMS, methanol, formate, and methylamine as described above.
All vials were sealed using sterile blue Teflon-coated butyl rubber septa. Filtered, sterile DMS solution was added aseptically through the septa of crimp-top vials with a syringe and needle to a final concentration of 50 µM from a 5 mM stock solution prepared with MAMS. Enrichment cultures preenriched on substrates other than DMS were later subcultured (10% inoculum) on DMS only (50 µM). Cultures were incubated at room temperature (20 to 25°C). Chemical controls consisting of medium supplemented with DMS were set up alongside enrichment cultures to account for chemical breakdown of DMS. The concentration of DMS in headspace gas was monitored by gas chromatography (GC) analysis. Enrichments were respiked with additional doses of DMS upon depletion of headspace DMS.
GC analysis.
Determination of DMS in headspace gas was carried out by injecting 100 µl of a headspace gas sample into a GCD gas chromatograph (PYE Unicam Ltd., Cambridge, United Kingdom) fitted with a 1 m-by-4 mm glass column containing Poropak Q (Phase Separations Ltd., Deeside, United Kingdom), and nitrogen as the carrier gas (flow rate, 30 ml min–1) at 200°C. A flame ionization detector was used to detect compounds, and peak areas were integrated with a model 3390A integrator (Hewlett Packard, Berkshire, United Kingdom). DMS concentrations were calculated by regression analysis based on a four-point calibration with standard DMS solutions in MAMS.
Isolation of bacterial strains and screening for DMS oxidation activity.
Samples of enrichment cultures were serially diluted in sterile MAMS medium, and 100 µl of sample was spread onto MAMS plates (MAMS solidified with 15 g liter–1 Bacto agar [Difco]). Plates were incubated for at least 2 weeks in gas-tight jars to which DMS was added (concentration of approximately 200 µM). Gas jars were regularly vented and replenished with DMS. Colonies were isolated and incubated as described above. Biomass from isolation plates was taken with a wire loop and resuspended in 1 ml of sterile MAMS and injected with sterile syringes through stoppers into 27-ml crimp-top vials containing 5 ml of sterile MAMS medium. DMS was added to a final concentration of 50 µM, and the degradation of DMS was monitored by GC analysis of headspace gas.
Test for growth on DMS.
Isolates were tested for their ability to grow on DMS on MAMS medium plates in gas-tight jars which contained DMS in the atmosphere (approximately 0.1% volume). To verify that isolates grew at the expense of DMS consumption and not on traces of organic compounds present in the solidified medium, degradation of DMS (50 µM) by isolates was also tested in liquid culture by monitoring headspace concentrations of DMS by gas chromatography. In addition, the growth of Methylophaga isolates was also tested at DMS concentrations of 500 µM and 1 mM. No growth was observed when Methylophaga strains were inoculated into medium lacking a carbon source. Isolates were also inoculated onto marine agar (2216; Difco) or into liquid MAMS medium to which peptone and yeast extract (44) were added, to test for the ability to grow on a complex medium.
PCR amplification of 16S rRNA-encoding genes, identification of isolates, and BOX-PCR of Methylophaga isolates.
Amplification and sequencing of bacterial 16S rRNA genes were done as described previously (44). For isolates, single colonies were taken from an agar plate with a sterile loop, resuspended in 50 µl of PCR-grade water, and boiled for 5 min. Lysates (1 to 5 µl) were used as the template for amplification of 16S rRNA genes by PCR, using primers 27F and 1492R (30). PCR products were obtained for all isolates, including gram-positive isolates. Two milliliters of enrichment cultures was pelleted at 13,000 x g at 4°C for 15 min in a microcentrifuge, and the pellet was resuspended in 10 µl of PCR-grade water and boiled for 5 min in a water bath. PCR products suitable for DGGE analysis were obtained as described previously, using primers 341F-GC and 926RM (45). Sequences were analyzed using BLAST (1) at the NCBI database (http://ncbi.nlm.nih.gov/BLAST) and added to those with the highest-scoring BLAST hits, to an alignment of bacterial 16S rRNA sequences (33) using the aligning tool included in ARB software (32). Phylogenetic trees were calculated using maximum-likelihood, parsimony, and distance methods. Bootstrap values were determined on 1,000 resampled data sets using PHYLIP programs SEQBOOT, DNADIST (with settings Kimura 2-parameter, transition/transversion ration of 2.0), NEIGHBOR, and CONSENSE (14). Genomic fingerprinting of Methylophaga isolates was carried out using BOX-PCR as described previously, using primer BOXA1R (42). The BOX-PCR method exploits conserved and repeated sequence motifs present in bacterial genomes that were first discovered in Streptococcus pneumoniae (35). Using the conserved sequence motif as a primer target site, a specific pattern of amplicons is generated that can be used for genomic fingerprinting of bacterial isolates (41).
DGGE and sequencing of DGGE bands.
DGGE was carried out as described previously (45), using gradients of 30 to 70% denaturants. DGGE staining with SYBR green I (Invitrogen, Paisley, United Kingdom) and image acquisition were carried out as described previously (39), using a FujiFilm FLA-5000 scanner. DGGE bands were sampled using sterile pipette tips and reamplified using primers 341F-GC and 926RM, as described previously (45). Bands were sequenced directly from purified PCR products using primer 926RM. If sequencing data were ambiguous due to mixed templates, PCR products were cloned using a TOPO-TA cloning kit (Invitrogen, Paisley, United Kingdom), and individual clones were reanalyzed by DGGE parallel to the original PCR product to identify comigrating clones, which were sequenced using standard M13 primers.
Effect of inhibitors on DMS metabolism by Methylophaga sp. strain DMS010.
An inhibition assay was carried out using biomass of strain DMS010 grown on DMS to an optical density (OD) (at 540 nm) of 0.3. Two hundred fifty milliliters of the culture was harvested by centrifugation at 17,700 x g at 15°C in a JA-10 rotor in a Beckman centrifuge. The cells were washed with sterile MAMS and resuspended in 25 ml of fresh medium. The assay was set up in triplicate in 27-ml crimp-top vials containing 5 ml of MAMS, 400 µl of a 3 mM DMS solution prepared in MAMS, 100 µl of inhibitor (50 mM chloroform or methyl tert-butyl ether [MTBE] in sterile distilled water or sterile water for controls, see below), and 500 µl of cell suspension (final optical density at 540 nm of approximately 0.3). Prior to the addition of cell suspension (or water for controls), the DMS-containing vials were left to equilibrate for 1 h. Uninoculated controls were set up in parallel to assess chemical losses of DMS.
Substrate-induced oxygen uptake of resting cell suspensions.
Methylophaga sp. strain DMS010 was grown in batch culture at 25°C in a shaking incubator at 150 rpm in 1.1-liter sealed crimp-top bottles in 250 ml MAMS medium and either 25 mM methanol or 1 mM DMS as the carbon source. Multiple cultures were grown on DMS and repeatedly respiked with DMS in order to obtain enough biomass for oxygen electrode experiments with DMS-grown cells. Cells were harvested by centrifugation at approximately 10,000 x g (15°C, 20 min) in a Beckman centrifuge using a JA-10 rotor and resuspended in 50 ml of sterile MAMS medium. The harvested cells were incubated for 2 h on a shaking incubator as described above before being used for the measurement of substrate-induced oxygen uptake rates, using a Clark-type oxygen electrode (Rank Brothers, Bottisham, United Kingdom) and a cell volume of 2 ml. The assay temperature was kept constant at 25°C by using a recirculating water bath. Substrates were added by using gas-tight syringes from concentrated stock solutions. Signals were recorded with a Philips PM8521A one-line recorder.
Analysis of polypeptides by SDS-PAGE, mass spectrometry, and N-terminal sequencing.
Methylophaga sp. strain DMS010 was grown on methanol (25 mM) and DMS (1 mM), and the biomass was harvested by centrifugation at 17,700 x g using a JA-10 rotor in a Beckman centrifuge at 4°C for 20 min. SDS-PAGE analysis of biomass from methanol- and DMS-grown Methylophaga sp. strain DMS010 was carried out using various percentages of acrylamide/bis-acrylamide as described previously (44). Polypeptide bands were excised from the gels and analyzed by mass spectrometry using matrix-assisted laser desorption ionization-mass spectrometry and in-line electrospray ionization tandem mass spectrometry at the Biological Mass Spectrometry and Proteomics Facility, Department of Biological Sciences, University of Warwick, as described previously (44). For N-terminal sequencing, SDS-PAGE gels were electroblotted onto polyvinylidene difluoride membrane (Amersham, United Kingdom) using a Novex Xcell blot module (Invitrogen) following the manufacturer's instructions. Blots were stained with Ponceau S (0.1% [wt/vol] in 1% [vol/vol] acetic acid), briefly rinsed in sterile deionized water, and air dried before target bands were cut out for N-terminal sequence analysis at Alta Bioscience (Birmingham, United Kingdom).
Nucleotide sequence accession numbers.
The nucleotide sequences obtained in this study have been deposited in the EMBL Nucleotide Sequence Database under accession numbers DQ660911 to DQ660973.
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RESULTS |
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PCR-DGGE analysis of DMS enrichment cultures and sequencing of DGGE bands.
DGGE analysis (Fig. 1) showed all enrichment cultures to be mixed cultures with common DGGE bands between enrichments obtained from the same sample. Several predominant bands were identical between DGGE profiles of enrichments obtained from different samples. E. huxleyi isolate-derived DMS-enrichment cultures had almost identical electrophoretic patterns, with a common dominant band observed for genetic fingerprints of all cultures. The DGGE profiles of samples from Achmelvich Bay and the rock pool that had been exposed to 500 µM DMS also had similar predominant bands.
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DMS-degrading enrichment cultures from Scottish coastal seawater samples, Achmelvich Bay, and the rock pool shared a number of phylotypes related to Methylophaga (Achmelvich, Table 1, band 15; rock pool, Table 1, bands 17 and 18), Alcanivorax (Achmelvich, Table 1, bands 15 and 25; rock pool, Table 1, band 20), and bacterial 16S rRNA genes identical to those of SCRIPPS_94, a sequence type identified in cultures of Scrippsiella sp. algae (Achmelvich, Table 1, band 26; rock pool, Table 1, band 21). In addition, one of the rock pool enrichments (50 µM DMS) contained a population related to an uncultured Actinomycetales bacterium (Table 1, band 22).
DGGE profiles of enrichments with samples from the English Channel (L4) had a higher number of bands than those from other samples. Affiliation of the sequences from dominant bands included Methylophaga (Table 1, band 30), a Gammaproteobacteria clade related to Methylophaga found in methane-rich environments (Table 1, band 50, enrichment with DMS, bicarbonate and thiosulfate; the best BLAST hit was clone HMMVCen-15, accession number AJ704664; T. Loesekann, T. Nadalig, H. Niemann, K. Knittel, A. Boetius, and R. Amann, unpublished data), Alcanivorax, members of the phyla Bacteroidetes and Firmicutes, members of the Roseobacter group, and Erythrobacter-like bacteria.
Isolation of bacterial strains from DMS enrichment cultures.
Twenty-four isolates were obtained from the enrichment cultures. These belonged to classes Alpha- and Gammaproteobacteria and to the Actinobacteria phylum. The identity of the isolates obtained by sequencing of 16S rRNA genes and the results of growth experiments with DMS are summarized in Table 2. The relationship of Methylophaga isolates to DGGE band sequences and other Methylophaga species is shown in Fig. 2. The sequences obtained from DGGE bands were all identical to those of Methylophaga strains isolated in this study, except for a few positions of sequence ambiguity. PCR products suitable for DGGE analysis obtained from Methylophaga isolates DMS002, DMS004, DMS009, and DMS010 comigrated with DGGE bands from enrichments cultures identified as Methylophaga populations (results not shown). Of the isolates that were obtained from DMS and methanol enrichment cultures, those related to the genera Methylophaga, Marinobacter, and Glaciecola were capable of oxidizing two consecutive additions of DMS (50 µM). Other cultures did not deplete the headspace of DMS. Growth of Methylophaga isolates DMS002, DMS004, DMS009, and DMS010 on DMS at concentrations of 500 µM and 1 mM was concomitant with an increase in optical density of liquid cultures (data not shown). Unlike the Methylophaga isolates, the Marinobacter and Glaciecola strains did not grow on DMS (50 µM or 500 µM). A number of cultures related to the Roseobacter clade were also tested for DMS oxidation. Leisingera methylohalidivorans strain MB2 has been reported previously to grow on DMS to a limited extent (34, 43); however, GC measurements of headspace concentrations of DMS in this study did not show any evidence of DMS degradation (50 µM). Other Roseobacter group isolates that were tested for DMS oxidation did not degrade DMS (50 µM) either, including the methyl halide-degrading strains 179, 198, and Roseovarius sp. strain 217 (44) and Ruegeria algicola (strain FF3), Roseovarius tolerans (DSM 11457), R. nubinhibens (DSM 15170), R. crassostreae (DSM16950), and R. mucosus (DSM 17069).
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Effect of inhibitors on DMS oxidation by Methylophaga sp. strain DMS010.
The effect of chloroform and of MTBE on DMS oxidation by Methylophaga sp. strain DMS010 was tested with DMS-grown cells; however, neither chloroform nor MTBE addition affected DMS oxidation rates compared to those of inhibitor-free controls (Fig. 3).
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SDS-PAGE analysis and identification of DMS-induced polypeptides in Methylophaga sp. strain DMS010.
Biomass of Methylophaga sp. strain DMS010 was obtained on methanol and DMS and polypeptides induced under the two growth conditions were analyzed by SDS-PAGE analysis of crude cell extracts (Fig. 4). A number of polypeptides appeared to be more highly expressed during growth on DMS than methanol-grown cells. The results of mass spectrometry analysis of excised polypeptide bands and N-terminal sequencing are reported in Table 3. The large subunit of methanol dehydrogenase (MxaF) was identified in biomass of cells grown on methanol and DMS. However, additional polypeptide bands were observed during growth on DMS (Fig. 4). These polypeptides included transketolase, a thiol-specific alkyl hydroxyperoxide reductase, a protein tentatively identified as a homolog of proteins predicted from the genome sequences of Silicibacter pomeroyi and Methylococcus capsulatus (Bath) as selenium binding proteins, and XoxF, a homolog of MxaF. The function of XoxF in methylotrophs is unknown (8).
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DISCUSSION |
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Diversity of DMS-degrading enrichment cultures.
DGGE analysis suggested that the dominant populations in many of the DMS-degrading enrichments were related to Methylophaga and Alcanivorax species. The identity of 16S rRNA gene sequences of Methylophaga DGGE bands and Methylophaga isolates from the same enrichments indicated that the strains represented the populations growing in the enrichment cultures. Some phylotypes (Fig. 1, bands 7 and 50) were present in enrichments that had as their closest relatives sequences obtained from environments characterized by the turnover of one-carbon substrates, e.g., a marine methanol-fed bioreactor (29) and the methane-rich sediments of the Haakon Mosby mud volcano (GenBank accession number AJ704664; Loesekann et al., unpublished data), suggesting their potential involvement in the turnover of DMS or intermediates of C1 metabolism. Other DGGE band sequences (e.g., Fig. 1, bands 2 and 6, clone 2, bands 16, 20, 21, 25, 26, 32, and 35) had the highest similarities to those of phylotypes detected in cultures of a variety of marine phytoplankton, especially those from dinoflagellates, which are key producers of DMSP, the precursor of DMS.
DMS degradation by isolated strains.
Of the 24 isolates that were obtained, only those identified as Methylophaga were able to grow on DMS, providing a clear link between the populations detected in enrichments and DMS degradation. Methylophaga is a genus of restricted and obligate methylotrophs (10-13, 21), i.e., obligate methylotrophic isolates that exclusively utilize C1 compounds and restricted methylotrophs that, in addition to C1 substrates, can utilize one or a few multicarbon compounds as growth substrates. Previously, Methylophaga sulfidovorans, isolated from a marine microbial mat, had been shown to degrade and grow on DMS (10). Recently, DGGE bands related to Methylophaga were also detected in marine DMS enrichments by Vila-Costa and colleagues (49); however, DMS-degrading Methylophaga isolates were not obtained in that study. Previously reported Methylophaga isolates were obtained from marine sediments or microbial mats (10, 22), and so the isolation of DMS-degrading Methylophaga strains from samples obtained from coastal water and seawater further offshore in this study demonstrates for the first time that certain Methylophaga species may also play a role in DMS oxidation in pelagic marine environments. In the current study, the presence of a Methylophaga sp. in nonaxenic cultures of E. huxleyi could indicate that Methylophaga may cooccur with E. huxleyi or other DMSP-producing phytoplankton in the environment. This is also suggested by the detection of Methylophaga sp.-related bacteria in marine mesocosms used to study bacterium-alga interactions (40) and in a culture containing the dinoflagellate Gymnodinium catenatum (GenBank accession number AY701420) (D. H. Green and C. J. S. Bolch, unpublished data). BOX-PCR demonstrated that the Methylophaga isolates obtained in this study represented at least four genetically different populations and suggested that the 16S rRNA gene sequences did not reflect the diversity at the strain level.
Other isolates that were obtained did not grow on DMS. While cell suspensions of Marinobacter and Glaciecola isolates degraded DMS (50 µM), DMS did not support the growth of these isolates. This may have been due to the utilization of DMS as a sulfur source or due to its conversion to DMSO. It is likely that additional DMS-degrading bacteria were present in these enrichments that could not be isolated with the culturing conditions used. This is concluded from the observation that DGGE analysis and sequencing of bands suggested that in some enrichments Methylophaga-related bacteria were not present.
Despite the potential of some members of the Roseobacter clade to transform organosulfur compounds such as DMSP, methanethiol, and DMS (6, 16, 38), growth on DMS is clearly not a common phenotype of Roseobacter clade bacteria. This is concluded from the observation that Leisingera methylohalidivorans, several Roseovarius isolates, Ruegeria algicola, Silicibacter pomeroyi, and the methyl halide-degrading strains 179, 198, and 217 (44), all members of the Roseobacter clade, were not able to grow on DMS. The observation that L. methylohalidivorans did not grow on DMS was similar to the findings by Schaefer and coworkers (43), who reported that the strain did not grow on 1.4 or 5 mM DMS but that it was able to increase in cell numbers on 50 µM DMS and was maintained over three subcultures. In the present study, L. methylohalidivorans did not degrade DMS, as determined by GC analysis of headspace gas, suggesting that the limited growth observed on DMS may previously have been due to trace organic constituents present in the medium or that the isolate had lost the ability to degrade DMS during serial transfer in the laboratory.
DMS metabolism of Methylophaga sp. strain DMS010.
Inhibition of DMS oxidation by MTBE and by chloroform has been used as a means to differentiate between the operation of the monooxygenase pathway and the methyltransferase pathway of DMS oxidation (20, 53). Neither MTBE nor chloroform had an inhibitory effect on DMS oxidation by strain DMS010. This was different from observations for Thiobacillus strains, in which a marked inhibition of DMS oxidation by MTBE was observed in Thiobacillus sp. strain T5, while chloroform strongly inhibited DMS oxidation by Thiobacillus sp. strain ASN-1 (53). However, strain DMS010 had a lower apparent Km (2.1 µM) for DMS than Thiobacillus sp. strain T5 (Ks of 90 µM), which opens up the possibility that the higher affinity for DMS in Methylophaga might preclude inhibition by either of the two inhibitors at relatively high DMS concentrations. Based on results with these inhibitors, a metabolic route of DMS oxidation in Methylophaga sp. strain DMS010 cannot be assigned, and so further studies of the biochemistry are essential. With Thiocapsa roseopersicina M11, aerobic DMS degradation was not inhibited by these compounds either (23). The Km for DMS of Methylophaga sp. strain DMS010 was comparable to those determined for Methylophaga sulfidovorans (Ks, 1.5 µM [10]), Thiocapsa roseopersicina M11 (Km, 2 µM [23]), and Hyphomicrobium strain EG (Ks, 3 µM [47]).
The induction of polypeptides during the growth of marine DMS-degrading isolates has not been studied previously. The role of some of the polypeptides detected in biomass of DMS-grown Methylophaga remains unknown in the absence of further genetic and biochemical data, but the peptides identified here are promising candidates for further study. The homolog of the large subunit of methanol dehydrogenase, XoxF, might have a role in the metabolism of DMS or in the degradation of the intermediate methanethiol; this role, however, will need to be investigated in future studies. Previously, mxaF' knockout mutants of Methylobacterium extorquens (similar to xoxF) were not affected in their ability to grow on methanol or methylamine, and a phenotype associated with this gene has not yet been identified (8). Induction of a thiol-specific alkyl hydroperoxide reductase during growth on DMS may be a consequence of thiol stress due to the production of methanethiol as an intermediate of DMS metabolism.
Conclusions and outlook.
The information presented here strongly suggests that Methylophaga spp. are involved in DMS degradation in seawater and therefore may be part of the population of marine methylotrophs that has been suggested to be responsible for this biogeochemical process (26). The strains of Methylophaga obtained in this study are the first DMS-degrading isolates of this genus obtained from seawater samples. Strain DMS010 differed in its DMS metabolism from that of Thiobacillus species and unidentified isolates based on inhibition assays (20, 53). Strain DMS010 had a low apparent Ks, indicating that it may be able to degrade DMS at typical environmental concentrations (1 to 5 nM) (25) or when DMS concentrations may reach high nM concentrations during the decay of phytoplankton blooms and even µM concentrations as observed in coral mucus (5). Degradation of DMS by bacteria in the upper mixed layer of the oceans is potentially carried out by diverse bacterial populations and metabolic pathways. Clearly, 16S rRNA gene sequences obtained by culture-independent means are of limited use to predict the potential of a given population to degrade DMS, since species closely related to DMS-degrading isolates may lack the potential to degrade DMS. Strains of some species (e.g., Rhodovulum sulfidophilum and Thiocapsa roseopersicina) may also be able to carry out DMS transformations by more than one pathway (23, 36). Studying the phylogenetic and functional diversity of DMS-degrading bacteria in the marine environment will require functional genetic markers that target key enzymes of DMS degradation pathways, such as DMS monooxygenase, methyltransferases, or other enzymes. The Methylophaga strains obtained in this study provide essential model organisms with which to analyze the metabolic pathways and biochemistry of DMS oxidation and to develop functional gene markers for studying the microbial ecology of marine DMS oxidation.
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ACKNOWLEDGMENTS |
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This work was supported by the Natural Environment Research Council with a postdoctoral fellowship (NE/B501404/1) and a project grant (NE/C001109/1).
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FOOTNOTES |
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Published ahead of print on 23 February 2007.
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-subclass of the class Proteobacteria. Appl. Environ. Microbiol. 65:3810-3819.
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