Editor: Julian Marchesi
Detection and isolation of chloromethane-degrading bacteria from the Arabidopsis thaliana phyllosphere, and characterization of chloromethane utilization genes
Article first published online: 8 JUN 2011
© 2011 Federation of European Microbiological Societies. Published by Blackwell Publishing Ltd. All rights reserved
FEMS Microbiology Ecology
Volume 77, Issue 2, pages 438–448, August 2011
How to Cite
Nadalig, T., Farhan Ul Haque, M., Roselli, S., Schaller, H., Bringel, F. and Vuilleumier, S. (2011), Detection and isolation of chloromethane-degrading bacteria from the Arabidopsis thaliana phyllosphere, and characterization of chloromethane utilization genes. FEMS Microbiology Ecology, 77: 438–448. doi: 10.1111/j.1574-6941.2011.01125.x
- Issue published online: 11 JUL 2011
- Article first published online: 8 JUN 2011
- Accepted manuscript online: 5 MAY 2011 07:40AM EST
- Received 24 February 2011; revised 23 April 2011; accepted 28 April 2011., Final version published online 8 June 2011.
- cmu genes;
- methyl halide metabolism
- Top of page
- Materials and methods
- Supporting Information
Chloromethane gas is produced naturally in the phyllosphere, the compartment defined as the aboveground parts of vegetation, which hosts a rich bacterial flora. Chloromethane may serve as a growth substrate for specialized aerobic methylotrophic bacteria, which have been isolated from soil and water environments, and use cmu genes for chloromethane utilization. Evidence for the presence of chloromethane-degrading bacteria on the leaf surfaces of Arabidopsis thaliana was obtained by specific quantitative PCR of the cmuA gene encoding the two-domain methyltransferase corrinoid protein of chloromethane dehalogenase. Bacterial strains were isolated on a solid mineral medium with chloromethane as the sole carbon source from liquid mineral medium enrichment cultures inoculated with leaves of A. thaliana. Restriction analysis-based genotyping of cmuA PCR products was used to evaluate the diversity of chloromethane-degrading bacteria during enrichment and after strain isolation. The isolates obtained, affiliated to the genus Hyphomicrobium based on their 16S rRNA gene sequence and the presence of characteristic hyphae, dehalogenate chloromethane, and grow in a liquid culture with chloromethane as the sole carbon and energy source. The cmu genes of these isolates were analysed using new PCR primers, and their sequences were compared with those of previously reported aerobic chloromethane-degrading strains. The three isolates featured a colinear cmuBCA gene arrangement similar to that of all previously characterized strains, except Methylobacterium extorquens CM4 of known genome sequence.
- Top of page
- Materials and methods
- Supporting Information
Chloromethane, the most abundant volatile halocarbon in the atmosphere (600 p.p.t.), may be responsible for about 15% of chlorine-catalysed ozone destruction in the stratosphere (Clerbaux et al., 2007). The global chloromethane budget is uncertain, with estimated known sources not fully accounting for identified sinks. The major sink for chloromethane likely involves reactions with hydroxyl radicals in the troposphere (Yoshida et al., 2004). Industrial emissions of the compound estimates range from <10% (Yoshida et al., 2004) to 60% (Trudinger et al., 2004) of total known emissions. Natural sources of chloromethane emissions include higher plants, grasslands, salt marshes, peatlands, wood-rotting fungi, senescent leaves, biomass burning and oceans, with vegetation possibly representing the major biotic source (Saito & Yokouchi, 2008). A methyltransferase responsible for chloromethane production was purified from leaves of Brassica oleracea (Attieh et al., 1995, 2000). In Arabidopsis thaliana, a protein encoded by the gene HOL (harmless to ozone layer) catalyses S-adenosyl-l-methionine-dependent methylation of chloride (Rhew et al., 2003; Nagatoshi & Nakamura, 2009). This enzyme is involved in the transformation of thiocyanate produced upon the degradation of glucosinolate, and its product methylisothiocyanate appears to play a role in the resistance of A. thaliana to bacterial infection (Nagatoshi & Nakamura, 2009). A physiological role for enzyme-produced chloromethane remains to be demonstrated.
Certain methylotrophic bacteria are able to use chloromethane as the sole carbon and energy source for growth (McDonald et al., 2002; Trotsenko & Doronina, 2003; reviewed in Schäfer et al., 2007; Table 1). Chloromethane-degrading bacteria have been isolated from diverse environments such as soils (Doronina et al., 1996; Miller et al., 1997; Coulter et al., 1999; McAnulla et al., 2001a), activated sludge (Hartmans et al., 1986; Traunecker et al., 1991; Freedman et al., 2004), freshwater (McAnulla et al., 2001a), and seawater (Schäfer et al., 2005), and include representatives affiliated to a wide range of genera including Aminobacter, Hyphomicrobium, Leisingera, Methylobacterium, Roseovarius (Alphaproteobacteria), Pseudomonas (Gammaproteobacteria) and Acetobacterium (Actinobacteria). However, the association of such bacteria with plants has not yet been investigated.
|Strains||Doubling time (h)||Specific activity (nmol min−1 mg−1protein)*|
|Methylobacterium extorquens CM4||7.3±1.1||26.1±5.8|
|Hyphomicrobium chloromethanicum CM2||4.9±0.3||24.8±2.2|
|Hyphomicrobium sp. strain MC1||5.1±0.3||29.5±4.0|
|Hyphomicrobium sp. strain AT2||5.9±0.7||28.0±0.3|
|Hyphomicrobium sp. strain AT3||18.1±0.5||22.8±2.0|
|Hyphomicrobium sp. strain AT4||19.8±0.5||21.6±0.2|
The only pathway for chloromethane catabolism by aerobic bacteria characterized so far was investigated in detail for strain Methylobacterium sp. CM4 (Doronina et al., 1996). This strain may now be affiliated to the species Methylobacterium extorquens on the basis of its complete genome sequence, including revised 16S rRNA gene sequences (GenBank NC_011757). A set of genes essential for growth on chloromethane (Vannelli et al., 1998, 1999), termed cmu genes for chloromethane utilization, was identified by minitransposon mutagenesis of strain CM4. These genes and the corresponding enzymes defined a specific pathway for chloromethane utilization in methylotrophic metabolism that depends on corrinoid and folate cofactors (Vannelli et al., 1999). The chloromethane dehalogenase enzyme, consisting of a corrinoid methyltransferase protein encoded by cmuA and of a tetrahydrofolate-dependent methyltransferase encoded by cmuB, was purified and characterized (Studer et al., 1999, 2001). The role in chloromethane utilization of cmuC, encoding another putative methyltransferase essential for chloromethane utilization (Vannelli et al., 1999), remains to be elucidated. Chloromethane-dependent expression of cmu genes was also demonstrated (Studer et al., 2002).
Several other chloromethane-degrading strains were subsequently reported (Woodall et al., 2001; McAnulla et al., 2001b; Schäfer et al., 2005; Warner et al., 2005) and also contained cmu genes. The cmuA gene has been used as a molecular biomarker of bacterial methyl halide metabolism in a large variety of environments (McAnulla et al., 2001a; Miller et al., 2004; Borodina et al., 2005; Schäfer et al., 2005). The plant environment, however, has not yet been explored in the context of bacterial chloromethane metabolism, despite the fact that some of the genera featuring representatives of chloromethane-degrading bacteria, notably Hyphomicrobium, Methylobacterium and Pseudomonas, are known to be efficient colonizers of the phyllosphere (Kinkel, 1997; Andrews & Harris, 2000; Trotsenko et al., 2001; Knief et al., 2008, 2010; Raja et al., 2008; Delmotte et al., 2009).
To investigate the occurrence of chloromethane-degrading bacteria in the phyllosphere, bacterial enrichments from leaves of A. thaliana, which is the best-characterized chloromethane-emitting plant at the molecular level (Rhew et al., 2003; Nagatoshi & Nakamura, 2009), were performed with the aim of isolating new bacterial strains able to grow with chloromethane as the sole carbon source, and to develop new molecular tools to efficiently detect and characterize gene signatures for chloromethane utilization in isolated strains and environmental DNA, in particular in the phyllosphere environment.
Materials and methods
- Top of page
- Materials and methods
- Supporting Information
DNA and bacterial strains
Strains M. extorquens CM4 and Hyphomicrobium chloromethanicum CM2, isolated in Russia from petrochemical factory soil (Doronina et al., 1996), and Hyphomicrobium sp. strain MC1, isolated from industrial sewage sludge (Hartmans et al., 1986), were laboratory stocks. Genomic DNA extraction of bacterial strains was performed using the Wizard Genomic DNA Purification Kit (Promega, Madison, WI). Total DNA of Aminobacter lissarensis CC495, Aminobacter ciceronei IMB-1, Rhodobacteraceae 179 and Rhodobacteraceae 198 was kindly provided by H. Schäfer (University of Warwick, UK).
Liquid cultures for enrichment and growth were performed in a mineral medium for methylotrophic bacteria (M3) containing (L-1 of distilled water) KH2PO4 (6.8 g), (NH4)2SO4 (0.2 g), NaOH (5 M) (5.85 mL), yielding a final pH of 7.2. After autoclaving, 1 mL L−1 medium each of calcium nitrate solution (25 g L−1) and of trace elements solution containing (mg L−1) FeSO4·7H2O (100), MnSO4·H2O (100), ZnSO4 (29.5), Co(NO3)2·6H2O (25), CuCl2·2H2O (25), Na2MoO4·2H2O (25), NH4VO3 (14.4), NiSO4·6H2O (10), H3BO3 (10) and 0.5 mL L−1 of H2SO4 (95%) were added.
Bacterial strains were cultivated and isolated on a solid medium containing (L-1 of distilled water) K2HPO4 (1.04 g), (NH4)2SO4 (0.2 g), NaH2PO4·H2O (0.65 g), MgSO4·7H2O (0.1 g), bromothymol blue (0.1 g) and agar (15 g) at a pH of 7.2. Calcium nitrate and trace elements were added after autoclaving as described for liquid M3 medium.
Enrichment culture and strain isolation from plant leaves
Enrichment cultures were set up by adding one leaf of greenhouse-grown A. thaliana Columbia (Col-0, ∼40 mg) to a 300-mL vial fitted with a sealed mininert valve cap (Sigma) and containing 50 mL of M3 medium. Following the addition of 12 mL (approximately 10 mM) chloromethane gas (Fluka, approximately 5 atm) to the headspace, enrichment cultures were incubated under shaking (100 r.p.m.) at 30 °C. After 24 h, leaf material was removed and 12 mL of chloromethane gas was again added. Enrichment cultures obtained after three successive subcultures were spread onto mineral agar plates and incubated in sealed, gas-tight jars containing 1.3% chloromethane (v/v). Chloromethane dehalogenation was indicated by the development of yellow colour on a green background around dehalogenating colonies, which were selected and purified on fresh solid M3 medium.
Resting cell suspensions were prepared from exponential-phase cultures (50 mL, OD600 nm<0.3), obtained with chloromethane as the sole carbon source. After centrifugation at 14 700 g for 10 min, cells were washed twice in 50 mM chloride-free phosphate buffer pH 7.0, and the cell pellet was resuspended in the same buffer (6 mL final volume). Protein determination was performed with 1 mL of cell suspension using the bicinchoninic acid assay and a commercial kit (Pierce). For activity measurements, 5 mL of cell suspensions of chloromethane-degrading strains were added to 17-mL Hungate vial capped with a gas-tight mininert valve (Sigma). Chloromethane gas (10 mL) was added in excess and the vial was incubated at 30 °C. At different times, aliquots (0.5 mL) were sampled through the valve with a 1-mL syringe, transferred to Eppendorf tubes on ice, centrifuged immediately, and the resulting supernatants were transferred to fresh Eppendorf tubes and kept frozen until further use. Control experiments with phosphate buffer and with cell suspensions of the nondechlorinating strain M. extorquens AM1 (Vuilleumier et al., 2009) were performed to evaluate the nonenzymatic degradation of chloromethane.
Chloride concentration was determined spectrophotometrically as [FeCl]2+ (λmax=340 nm) formed in a highly acidic medium using the method of Jörg & Bertau (2004). Chloride concentration was determined by comparison with a calibration curve (0–5 mM) obtained with a sodium chloride solution in the same buffer, and dehalogenase activity was expressed as nmol min−1 mg−1 protein.
DNA extraction from bacterial cultures and environmental samples
Genomic DNA extraction from enrichment cultures (10 mL at OD600 nm=0.6) was performed using the Wizard Genomic DNA Purification Kit (Promega) according to the manufacturer's recommendations. DNA was extracted from epiphytic bacteria on the surface of A. thaliana leaves. Briefly, 10–15 leaves of A. thaliana plants (360–860 mg fresh weight) were washed as described previously (Delmotte et al., 2009), with 30 mL TE buffer, pH 7.5, containing 0.1% Silwet L-77 (GE Bayer Silicones). After centrifugation, total DNA was prepared using the FastDNA spin kit (MP Biomedicals, Santa Ana, CA), as described by Knief et al. (2008). Cell lysis was performed using a Mikro-Dismembrator S (Sartorius Stedim Biotech, France) by three consecutive 1-min treatments at 3000 min-1.
Quantitative PCR (qPCR)
CmuA and 16S rRNA gene copy numbers were evaluated through qPCR using an ABI PRISM 5700 sequence detection system (Applied Biosystem, Foster City, CA). qPCR analysis was carried out in triplicate using phyllospheric DNA (5–10 ng) in 96-well plates, using the primer pairs cmuA802f and MF2 (5′-CCRCCRTTRTAVCCVACYTC) for the cmuA gene and BACT1369F and PROK1492R (Suzuki et al., 2000) for the 16S rRNA gene, respectively. The PCR reaction mix contained 1 × qPCR Mastermix Plus for SYBR Green I (Eurogentec S.A., Belgium), 0.3 and 12.8 μM of cmuA802f and MF2 primers, respectively, for cmuA amplification, or 0.4 μM of each primer BACT1369F and PROK1492R for 16S rRNA gene amplification. Reaction conditions were 10 min at 95 °C, followed by 45 cycles of denaturation at 95 °C for 15 s, followed by annealing and elongation at 60 °C for 60 s. Calibration curves were obtained using genomic DNA from M. extorquens CM4 for both cmuA and 16S rRNA gene analysis.
PCR and RFLP analysis
The primers used in this study are listed in Table 2. Reactions were performed in 0.2-mL microcentrifuge tube using a thermal cycler (Mastercycler Personal, Eppendorf, Germany). Each PCR reaction mixture consisted of 2.5 μL of PCR buffer (New England Biolabs), 0.25 μL of dNTPs (200 μM), 1 μL of each forward and reverse primers (20 μM), 17.9 μL of distilled water, 0.3 μL of Taq polymerase (5 U μL−1, New England Biolabs), 0.05 μL of Pfu polymerase (3 U, Promega) and 2 μL of template DNA solution (25 ng). After initial denaturation (94 °C, 3 min), DNA amplification was performed by 30 cycles of 45-s denaturation at 94 °C, annealing for 1 min (at 52 °C for the 16S rRNA gene and between 61 and 67 °C for cmu genes; see Table 2 for details), extension for 1–4.5 min (depending of fragment length) at 72 °C and a final extension step of 7 min at 72 °C.
|Target gene||Primer*||Sequence (5′–3′)†||Position‡||References|
|cmuA||cmuA802f||TTCAACGGCGAYATGTATCCYGG||7404–7426||Miller et al. (2004)|
|cmuA1609R||TCTCGATGAACTGCTCRGGCT||8212–8190||Miller et al. (2004)|
|hutI||hutIrev2||TCVTCRCARHAVRCYTCDAC||10 655–10 635||This study|
A two-step semi-degenerate PCR strategy (Jacobs et al., 2003) was used to access the 5′-upstream region of the cmuB gene fragments obtained. In the first PCR, primer cmuBrev was used with a mix of the three semi-degenerate primers cekg2A, cekg2B and cekg2C (Jacobs et al., 2003). The second PCR involved using the reverse primer cmuBrev2 and the primer cekg4 targeting the tail of the semi-degenerate primers used in the first PCR (Jacobs et al., 2003).
Amplified cmuA fragments from each strain and from enrichment cultures were purified using the ‘GENECLEAN Turbo’ kit (MP Biomedicals Europe, France). Purified products (2 μg) were digested with the enzyme DdeI (30 U; Fermentas, France) and the appropriate buffer at 37 °C for 20 h. Digested products were size fractionated on 2% agarose/Nusieve (3/1) gels.
PCR products were purified with ExoSAP-IT reagent (USB Corporation) according to the manufacturer's recommendations; DNA sequences were obtained from PCR products with appropriate primers (Table 2) on an ABI Prism 3130 XL Genetic Analyzer (Applied Biosystems, UK). The assembled sequences for the cmuBCA clusters of Hyphomicrobium strains MC1, AT2, AT3 and AT4 obtained were deposited in the EMBL database under numbers FN667867, FN667868, FN667869 and FN667870 respectively, together with the corresponding partial rRNA gene sequences (FN667863, FN667864, FN667865 and FN667866, respectively).
DNA sequences were compared with databases by online blast searches (http://www.ebi.ac.uk/tools/blast). DNA and protein sequences of interest were analysed using the Mobyle platform (http://mobyle.pasteur.fr). Multiple alignments were obtained using clustalw, and manually adjusted using jalview, as implemented on the Myhits online portal (http://myhits.isb-sib.ch). Multiple alignments were analysed using the phylip suite of programs, and in particular the dnadist, neighbor, seqboot and consense, as implemented on the Mobyle platform. Bootstrap analysis was performed with 100 replicates.
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- Materials and methods
- Supporting Information
Quantification of 16S rRNA and cmuA genes on the leaf surfaces of A. thaliana
Total DNA was extracted from the leaf surface of leaves A. thaliana, yielding approximately 0.4–1.2 ng DNA mg−1 fresh weight of leaves (range 30–1000 ng DNA per leaf). The cmuA gene was detected in all samples, with a copy number of 8.0±0.8 copies ng−1 DNA. The 16S rRNA gene was detected in the same samples at a copy number of 10 100±2000 copies ng−1 DNA. Assuming a single copy of 16S rRNA gene per bacterial genome, this suggested that on average, <1 in 1000 bacterial cells on leaf surfaces carried the cmuA gene and may be capable of dehalogenating chloromethane.
Isolation, characterization and identification of chloromethane-utilizing bacteria
Enrichment cultures were set up with A. thaliana leaves as the inoculum in a chloride-free mineral medium with chloromethane as the sole carbon and energy source (1.3%, v/v). Chloromethane-dependent growth with concomitant chloride production was observed in enrichment cultures. Three chloromethane-degrading bacterial strains termed AT2, AT3 and AT4 (AT in reference to A. thaliana) were obtained from such enrichment cultures, as single colonies on a solid mineral medium with chloromethane as the sole carbon and energy source. These isolates grew aerobically on both liquid and solid mineral medium with chloromethane, methanol or succinate as the sole carbon source, indicating that they were facultative methylotrophs. All three strains displayed characteristic hyphae, indicative of the genus Hyphomicrobium (Moore, 1981). Taxonomical affiliation to cluster II (Rainey et al., 1998) was confirmed using 16S rRNA gene sequence analysis (Fig. 1). The ability of strains AT2, AT3 and AT4 to transform chloromethane and to use it as the sole carbon and energy source for growth was compared with reference strains M. extorquens CM4 and H. chloromethanicum CM2 (Table 1). Hyphomicrobium strains AT2 grew with chloromethane with similar doubling times as the previously described strains CM2 and MC1 (td∼5 h). The growth of M. extorquens CM4 was slightly slower (td 7.3 h), whereas newly isolated Hyphomicrobium strains AT3 and AT4 were the slowest growing (td over than 18 h). In contrast, specific chloromethane dehalogenation activities inferred from measurements of chloride concentration in the supernatants of cell suspensions were similar for all strains (Table 1).
Organization and diversity of cmu genes in chloromethane-degrading strains
Genes cmuA, cmuB and cmuC could be amplified and sequenced from total DNA of chloromethane-degrading phyllosphere isolates using both previously described primers (McAnulla et al., 2001b; Miller et al., 2004) and primers newly designed from the better conserved sequence regions in previously reported cmu gene clusters (Table 2). A two-step PCR strategy (Jacobs et al., 2003) afforded access to the unknown sequence region upstream of amplified cmuB gene fragments yielding complete sequences of the cmuB gene for the phyllosphere isolates. The sequences and organization of cmu genes in strains isolated from A. thaliana were compared with those of previously described strains (Figs 2 and 3, Supporting Information, Fig. S1).
Phylogenetic analysis of partial cmuA gene sequences from phyllosphere isolates obtained using the primer pair cmuA802f-cmuA1609R (Miller et al., 2004) was compared with those of previously reported chloromethane-degrading strains, and from selected cmuA gene fragments available in sequence databases and obtained from environmental DNA of different origins (Miller et al., 2004; Borodina et al., 2005; Schäfer et al., 2005) (Fig. 3). This analysis yielded a picture congruent with that obtained for the analysis of the 16S rRNA gene (Fig. 1). It also suggested that phyllosphere isolates, together with strain MC1, belong to a clade that includes sequences from woodland soil covered with leaf-litter and garden soils (Borodina et al., 2005). The levels of sequence identity between cmuA gene fragments were 75–80% between Hyphomicrobium strains and either M. extorquens CM4 or Aminobacter strains. cmuA amplicons of strains AT3 and AT4 showed identical sequences, differing from those of strains AT2, CM2 and MC1 which, with over 99% pairwise identity, clustered tightly together (Fig. 3).
In contrast with cmuA, only a few partial or full cmuB and cmuC gene sequences are available so far, all obtained from cultivated and isolated strains (Fig. 2). The new degenerate primer pairs developed in this work allowed the detection and retrieval of cmuB and cmuC gene sequences from the newly isolated chloromethane-degrading strains reported here, and comparison with previously described cmu genes (Table 2). Overall, the sequences for cmuB gene fragments (Fig. S2a) showed high levels of identity, but cmuB sequences of Hyphomicrobium strains were only about 60% identical to that of M. extorquens CM4. Regarding cmuC (Fig. S2b), sequence analysis of amplicons again showed that sequences from AT3 and AT4 were most closely related (97.9% identity), and that sequences from strains MC1, AT2 and CM2 formed a closely related cluster (>91% identity). Notably, cmuC sequences of Hyphomicrobium strains including the new isolates were equally distant (∼47% identity) to cmuC and to cmuC2 of unknown function found immediately upstream of cmuA, of strain CM4. This emphasizes the lesser degree of conservation of cmuC despite it being essential for growth with chloromethane in strain CM4 (Vannelli et al., 1999).
The cmuBCA cluster organization of cmu genes for Hyphomicrobium strains isolated from A. thaliana leaves and for strain MC1 was the same as that found previously for H. chloromethanicum CM2 and all other previously isolated strains, with the exception of M. extorquens CM4 (Fig. 2). In all cases where it was characterized, this single-cluster cmu gene arrangement also featured genes paaE and hutI, encoding a putative oxidoreductase and a putative imidazolone hydrolase, respectively. In this work, PCR reactions with cmuA802f-paaErev1 and cmuA802f-hutIrev2 primer pairs (Table 2, and data not shown) provided evidence that paaE and hutI genes were also present in the three new strains reported here and in the same arrangement as in strain H. chloromethanicum CM2 (McAnulla et al., 2001b).
New PCR primers for cmuA analysis
The published reverse primer cmuA1609R used for the detection of cmuA sequences in environmental samples (Miller et al., 2004) allowed to accommodate the shorter cmuA sequence of A. ciceronei. In this work, a frameshift in the cmuA sequence originally reported for A. ciceronei IMB1 (AF307143) sequence was detected and corrected, extending its predicted cmuA gene from 1704 to 1851 nt. Amplification using the primer pair cmuA802f together with cmuA1802r newly designed in this work yields a larger cmuA gene PCR fragment of approximately 1 kb (Table 2). Similar sensitivity was achieved with the newly designed cmuA primers and with the previously described primers (see Fig. S1). However, the new primer cmuA1802r may allow the detection of a wider diversity of cmuA sequences, because it could be successfully used to amplify cmuA from marine strain Rhodobacteraceae 198, unlike primer cmuA1609R (Schäfer et al., 2005) (Fig. 4). Also, the 193-bp 3′-end cmuA sequence also amplified with this newly defined primer pair is slightly less conserved than the sequence between primers cmuA802f and cmuA1609R (see Table S1), thus potentially allowing better discrimination of cmuA sequences retrieved from environmental DNA.
Restriction fragment profiling of PCR-amplified cmuA in chloromethane-degrading enrichment cultures obtained from plant leaves
A protocol for monitoring chloromethane-degrading enrichment cultures obtained from leaves of A. thaliana as inocula was developed using restriction digestion of PCR-amplified cmuA gene fragments (Fig. 4). At the timepoint chosen to isolate chloromethane-degrading strains by plating out of the liquid enrichment culture on a solid selective mineral medium (OD600 nm=0.6, 8 days), the detection limits using primer pair cmuA802f-cmuA1802r developed here were typically 0.5 and 10 pg of the DNA template for the reference strain M. extorquens CM4 and for the enrichment culture, respectively (Fig. S1). This suggested that the chloromethane-degrading bacterial subpopulation in enrichment cultures represented about 5% of the total bacteria present in the cultures at that stage.
The amplicons obtained were digested with the restriction enzyme DdeI, which cuts one to three times and at variable positions in the cmuA sequence of previously characterized chloromethane-degrading strains (Schäfer et al., 2005) (Fig. 4). The digestion patterns of amplicons in enrichment cultures were distinct from all reference strains, except for H. chloromethanicum CM2 and Hyphomicrobium sp. MC1. Strain AT2 showed the same pattern as the enrichment culture from which it was isolated (Fig. 4), and the same situation was found for strains AT3 and AT4 and the corresponding enrichment culture from which these strains originated (data not shown). The restriction profiling method applied here to amplicons of 1000 nucleotides thus holds promise for a focused, time-saving exploration of chloromethane utilization genes aiming at discovering new, more divergent cmu gene sequences in environmental samples, and at characterizing the corresponding bacteria.
Bacteria growing aerobically with chloromethane as the sole source of carbon and energy had previously been isolated from a variety of environments, and so far, all feature the cmu pathway for chloromethane utilization (Studer et al., 2002; Schäfer et al., 2007). However, the phyllosphere compartment of vegetation, possibly the quantitatively most important source of chloromethane (Clerbaux et al., 2007), had not yet been investigated in this respect. In this work, the key gene cmuA involved in dehalogenation of chloromethane was detected and quantified in DNA from the leaf surface of the model plant A. thaliana, and three chloromethane-degrading Hyphomicrobium strains were isolated from enrichment cultures originating from leaves of A. thaliana grown with chloromethane as the sole carbon source. In addition, several degenerate primer pairs and an associated genotyping approach were developed for the detection and characterization of cmu genes in enrichment cultures and isolated strains.
The Hyphomicrobium chloromethane-degrading strains isolated in this work possessed cmu genes in the same arrangement as in most previously isolated strains from other ecosystems, confirming the dominant status of the cmu pathway in the bacterial degradation of chloromethane. However, the isolation of strains belonging to the Hyphomicrobium genus was unexpected, inasmuch as Methylobacterium strains were recently shown to be efficient leaf colonizers and predominant in the A. thaliana phyllosphere, with Hyphomicrobium likely representing only a minor contribution (Delmotte et al., 2009; Knief et al., 2010). Indeed, enrichment cultures obtained from plant leaves in the same medium, but with methanol as the sole carbon and energy source led to the enrichment of strains belonging to the genus Methylobacterium (data not shown). However, chloromethane-degrading Hyphomicrobium isolates also grew well with methanol as the carbon source, suggesting that Hyphomicrobium strains may be better adapted to growth with chloromethane than Methylobacterium strains, albeit in an as yet unknown way. Other aspects of Hyphomicrobium metabolism require further investigation, in particular, the fact that similar chloromethane dehalogenase activity was detected in cell-free extracts of all chloromethane-degrading strains despite differences in the growth rates of the strains with chloromethane (Table 1).
The demonstration of chloromethane-degrading bacteria at the surface of A. thaliana leaves is of relevance for the overall budget of chloromethane in the environment in the light of current estimates for chloromethane emissions above plant areas (∼1.8 Tg Cl year−1) (Yoshida et al., 2006). If indeed, as suggested from this work, some phyllosphere bacteria function as a filter for emissions of chloromethane from plants, then measurements and estimates of chloromethane emissions above plant areas will actually tend to reflect the difference between total chloromethane emissions from vegetation and bacterial degradation of chloromethane in the phyllosphere, rather than the total chloromethane potential from plants. Whether this may contribute towards explaining the deficit in identified sources of chloromethane (Clerbaux et al., 2007) is a topic for further investigation. Clearly, assessing the importance of chloromethane degradation by specialized methylotrophic bacteria in the phyllosphere will require further work, especially considering that plant emissions of methanol, itself a growth substrate for most methylotrophic bacteria, exceed those of chloromethane by over three orders of magnitude (Nemecek-Marshall et al., 1995; Rhew et al., 2003).
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- Materials and methods
- Supporting Information
Support for this project by REALISE, the Alsace Network for Engineering and Environmental Sciences (http://realise.u-strasbg.fr) and from the EC2CO program of CNRS-INSU is gratefully acknowledged. We also thank SFERE, the Government of Pakistan and the French Ministry of Foreign Affairs for a PhD grant to M.F.U.H., and the French Ministry of Research and Higher Education for a PhD grant to S.R.
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- Materials and methods
- Supporting Information
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- Top of page
- Materials and methods
- Supporting Information
Fig. S1. Analysis of PCR-amplified fragments of cmuA obtained with primers cmuA802f and cmuA1802r.
Fig. S2. Phylogenetic analysis of cmuB and cmuC genes in chloromethane-degrading strains.
Table S1. Sequence identity of amplified cmuA gene fragments.
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