Chloromethane-Dependent Expression of the cmu Gene Cluster of Hyphomicrobium chloromethanicum

Abstract

The methylotrophic bacterium Hyphomicrobium chloromethanicum CM2 can utilize chloromethane (CH₃Cl) as the sole carbon and energy source. Previously genes cmuB, cmuC, cmuA, and folD were shown to be essential for the growth of Methylobacterium chloromethanicum on CH₃Cl. These CH₃Cl-specific genes were subsequently detected in H. chloromethanicum. Transposon and marker exchange mutagenesis studies were carried out to identify the genes essential for CH₃Cl metabolism in H. chloromethanicum. New developments in genetic manipulation of Hyphomicrobium are presented in this study. An electroporation protocol has been optimized and successfully applied for transformation of mutagenesis plasmids into H. chloromethanicum to generate stable CH₃Cl-negative mutants. Both transposon and marker exchange mutageneses were highly applicable for genetic analysis of Hyphomicrobium. A reliable and reproducible selection procedure for screening of CH₃Cl utilization-negative mutants has also been developed. Mutational inactivation of cmuB, cmuC, or hutI resulted in strains that were unable to utilize CH₃Cl or to express the CH₃Cl-dependent polypeptide CmuA. Reverse transcription-PCR analysis indicated that cmuB, cmuC, cmuA, fmdB, paaE, hutI, and metF formed a single cmuBCA-metF operon and were coregulated and coexpressed in H. chloromethanicum. This finding led to the conclusion that, in cmuB and cmuC mutants, impaired expression of cmuA was likely to be due to a polar effect of the defective gene (cmuB or cmuC) located upstream (5′) of cmuA. The detrimental effect of mutation in hutI on the upstream (5′)-located cmuA is not clear but indicated that all the genes located within the cmuBCA-metF operon are coordinately expressed. Expression of the cmuBCA-metF transcript was also shown to be strictly CH₃Cl inducible and was not repressed by the alternative C₁ substrate methanol. Sequence analysis of a transposon mutant (D20) led to the discovery of the previously undetected hutI and metF genes located 3′ of the paaE gene in H. chloromethanicum. MetF, a putative methylene-tetrahydrofolate reductase, had 27% identity to MetF from M. chloromethanicum. Mutational and transcriptional analysis data indicated that, in H. chloromethanicum, CH₃Cl is metabolized via a corrinoid-specific (cmuA) and tetrahydrofolate-dependent (metF, purU, folD) methyltransfer system.

Chloromethane (CH₃Cl), with an average atmospheric concentration of 500 to 600 pptv (parts per trillion by volume), represents the most abundant halocarbon in the atmosphere (17, 49). CH₃Cl is mainly of natural origin and is responsible for about 17% of chlorine-catalyzed ozone destruction (18, 19). Current sources of CH₃Cl that have been identified include biomass burning, emissions from oceans, salt marshes, wood-rotting fungi, higher plants, coal combustion, and industrial emissions (23, 24, 27, 36, 38, 52, 55). The dominant loss process for CH₃Cl is via reaction with OH radicals in the atmosphere, but soils have also been identified as a significant sink for CH₃Cl, where it is mainly degraded by methylotrophic bacteria (6, 9, 20, 25, 30, 33, 34, 49).

The four extant soil CH₃Cl utilizers, Methylobacterium chloromethanicum, Hyphomicrobium chloromethanicum, and Aminobacter sp. strains IMB-1 and CC495, have been examined at the physiological and molecular level to investigate the mechanism of CH₃Cl metabolism (6, 31, 41, 44, 50, 51, 54). H. chloromethanicum and Aminobacter strains IMB-1 and CC495 also grow on bromomethane (CH₃Br). Leisingera methylohalidivorans, the only marine isolate, is able to utilize CH₃Cl, CH₃Br, and iodomethane (CH₃I) but has not been characterized with regard to its pathway of CH₃Cl oxidation (40). L. methylohalidivorans (40) and a soil isolate, Aminobacter sp. strain IMB-1 (41) were shown to be able to oxidize tropospheric concentrations of CH₃Br (12 pptv), indicating that these and other methyl halide utilizers are responsible for the biological oxidation of tropospheric CH₃Br and possibly CH₃Cl in various terrestrial and marine environments (15). This also indicated that ambient concentrations of CH₃Br and possibly CH₃Cl were adequate for the induction of methyl halide utilization pathway(s).

All four CH₃Cl-utilizing soil isolates were found to possess the gene cmuA along with other CH₃Cl-specific genes within their cmu (chloromethane utilization) gene clusters (6, 31, 32, 50, 54). The cmuA gene encodes methyltransferase I (CmuA), which catalyzes the initial dehalogenation step of the CH₃Cl degradation pathway (Fig. 1). CmuA is a 67-kDa polypeptide which is induced in a strictly CH₃Cl- and CH₃Br-dependent manner. Moreover, the CmuA polypeptide uniquely consists of two domains, a methyltransferase domain and a corrinoid-binding domain (6, 31, 45, 51, 54). This feature was exploited for designing PCR primer sets which amplify a fragment of cmuA spanning both domains. These PCR primers have been used successfully as a functional gene probe to detect cmuA in extant CH₃Cl utilizers, in enrichment cultures, and also in DNA extracted from natural soil and marine environments (30, 31, 32, 35).

FIG. 1.

Proposed pathway for metabolism of CH₃Cl in M. chloromethanicum CM4. The figure is adapted from Vannelli et al. (50). The techniques used to generate CH₃Cl-negative mutants of M. chloromethanicum are shown on the left. Enzymes involved in CH₃Cl metabolism in M. chloromethanicum are: CmuA, methyltransferase I; CmuB, methyltransferase II; MetF, putative 5,10-methylene-H₄-folate reductase; FolD, putative 5,10-methylene-H₄-folate dehydrogenase/5,10-methenyl-H₄-folate cyclohydrolase; PurU, putative 10-formyl-H₄-folate hydrolase; FDH, formate dehydrogenase.

The CH₃Cl utilization pathway (Fig. 1) was initially postulated on the basis of a transposon mutagenesis study carried out to create mutants of Methylobacterium chloromethanicum that did not grow on CH₃Cl (44, 50, 51). These molecular genetic studies demonstrated that both cmuA and cmuB (encoding methyltransferase II) are involved in the dehalogenation of CH₃Cl in M. chloromethanicum. Dehalogenation is initiated by methyl group transfer from CH₃Cl onto a vitamin B₁₂ cofactor (corrinoid group) bound to the protein CmuA (methyltransferase I) (45). The methyl group is sequentially transferred from CmuA to methyltetrahydrofolate (methyl-H₄-folate) by CmuB (methyltransferase II) (45, 46). Methyl-H₄-folate is then further oxidized to CO₂, via formate, to provide reducing equivalents for biosynthesis and energy generation (Fig. 1). Carbon is assimilated at the level of methylene tetrahydrofolate, directly feeding into the serine pathway.

Mutagenesis studies on M. chloromethanicum revealed that transposon insertion in cmuA, cmuB, cmuC, or purU caused loss of the ability to grow on CH₃Cl, indicating the essential role of all four of these genes for the metabolism of CH₃Cl. Sequencing of the genes containing transposon insertions mapped the CH₃Cl utilization genes to two separate clusters. cmuA, purU and folD are located within the same CH₃Cl utilization gene (cmu) cluster in M. chloromethanicum. Genes metF, cmuB, and cmuC are located on a separate cmu cluster in M. chloromethanicum and were recently reported to be cotranscribed in M. chloromethanicum (45, 46). An insertional mutagenesis study of metF in M. chloromethanicum revealed the essential role of this gene in CH₃Cl utilization. The mutagenesis studies on M. chloromethanicum suggested a novel CH₃Cl-specific C₁ utilization pathway, involving metF, folD, and purU, for the formation of formate from methyl-H₄-folate.

This H₄-folate-dependent CH₃Cl pathway, which is strictly induced by CH₃Cl in M. chloromethanicum, is thought to operate in all four extant CH₃Cl utilizers, which were found to possess CH₃Cl-specific genes. H. chloromethanicum and Aminobacter sp. strain IMB-1, possessing single cmu clusters, have an identical arrangement of cmuC, cmuA, fmdB, and paaE (31, 32, 54). cmuC is located downstream (3′) of cmuB in H. chloromethanicum, and paaE is found upstream (5′) of the hutI and metF genes in Aminobacter sp. strain IMB-1. Although folD is present at the 5′ end of the cmu cluster in H. chloromethanicum, the metF gene has not yet been identified (31). Detection and subsequent mutagenesis of metF in the genome of H. chloromethanicum would strongly support the proposed functioning of the H₄-folate-dependent CH₃Cl utilization pathway in H. chloromethanicum.

The novel H₄-folate-dependent CH₃Cl utilization pathway (Fig. 1) was detected and analyzed in M. chloromethanicum, but the presence of this pathway in H. chloromethanicum and Aminobacter strains IMB-1 and CC495 is purely speculative at present. Moreover, in Aminobacter sp. strain CC495, the involvement of halomethane:bisulfide/halide ion methyltransferase in CH₃Cl metabolism has been proposed, which possibly catalyzes methyltransfer from CH₃Cl or CH₃Br to an acceptor ion (such as HS⁻) to form methanethiol (6). As in M. chloromethanicum, H. chloromethanicum, and Aminobacter sp. strain IMB-1, the methyltransferase CmuA detected in Aminobacter sp. strain CC495 was found to be a 67-kDa protein containing a corrinoid group.

We initiated molecular genetic studies to explore the mechanism of CH₃Cl metabolism in H. chloromethanicum by applying transposon mutagenesis, marker exchange mutagenesis, and reverse transcription (RT)-PCR analysis. Here we report the analysis of mutants of H. chloromethanicum defective in CH₃Cl utilization. Sequence analysis of one of the transposon mutants provided information on an extended cmu cluster of H. chloromethanicum, containing hutI and metF, which were cloned, sequenced, and analyzed. Mutagenesis and transcriptional analysis provided evidence for the functioning of the H₄-folate-dependent CH₃Cl utilization pathway in H. chloromethanicum. In addition, an electroporation protocol for H. chloromethanicum was optimized and efficiently applied as a DNA transfer method to generate CH₃Cl utilization-negative mutants. Transcriptional analysis of the cmu cluster of H. chloromethanicum is also presented.

MATERIALS AND METHODS

Growth media and strains.

H. chloromethanicum strain CM2^T was routinely cultured at 30°C with shaking (200 rpm.) in 25-ml batch cultures on Paracoccus versutus medium (pH 7.4) (4). For growth on CH₃Cl, 125-ml serum vials sealed with Teflon-coated butyl rubber stoppers (Owens Polyscience Ltd., Macclesfield, United Kingdom) were used. CH₃Cl was added directly to the headspace through rubber stoppers to a final concentration of 2 or 5% (vol/vol) in the gas phase unless otherwise stated. When used, methanol was added to a concentration of 0.5% (vol/vol) in the liquid phase. Plate cultures of H. chloromethanicum were grown on P. versutus agar in sealed gas-tight jars gassed with 2% (vol/vol) CH₃Cl and incubated at 30°C. Escherichia coli strains TOP10 and CC118(λpir) (29) were grown in Luria-Bertani medium at 37°C. Antibiotics for E. coli and H. chloromethanicum were used at the following final concentrations: kanamycin at 50 μg ml⁻¹ and gentamicin at 5 μg ml⁻¹.

Determination of CH₃Cl by gas chromatography.

Samples of headspace gas (100 μl) were injected into a GCD gas chromatograph (Pye Unicam Ltd., Cambridge, United Kingdom) fitted with a Porapak Q column (Phase Separation Ltd., Deeside, United Kingdom) at 200°C. A flame ionization detector was used to detect products, and the peak areas were determined with a 3390A integrator (Hewlett-Packard, Berkshire, United Kingdom). The gas chromatograph was calibrated with samples of the headspace gases above standard solutions containing known concentrations of gases equilibrated at 30°C.

Gene transfer experiments. (i) Chemical transformation.

Competent cells of E. coli CC118(λpir) (21) for chemical transformations were prepared by washing with ice-cold 0.1 M CaCl₂ according to the method of Hanahan (16). Competent cells of E. coli TOP10 were purchased from Invitrogen and used according to the manufacturer's instructions.

(ii) Electroporation.

Electrocompetent cells of H. chloromethanicum were prepared by harvesting chilled cultures (400 ml) in early exponential phase at an optical density at 540 nm (OD₅₄₀) of 0.3 and carefully washing in ice-cold water (4,000 × g, 15 min, 4°C), followed by the same wash in ice-cold 10% (vol/vol) glycerol and then resuspension in 800 μl of 10% (vol/vol) glycerol; 50-μl aliquots of cells were mixed with 500 ng of plasmid DNA and incubated on ice for 5 min prior to electroporation. Electroporation was carried out in 0.1-cm gap cuvettes (Bio-Rad) with a Bio-Rad gene pulser (Bio-Rad Laboratories, Hercules, Calif.) with the following electrical settings: 2.4 kV and 200 Ω at a capacitance of 25 μF. Immediately after electroporation, 1 ml of P. versutus medium containing 0.5% (vol/vol) methanol was added to the cuvette, and cells were transferred to an Eppendorf tube and incubated with shaking for 1 h at 30°C. Transformants were selected by plating suitable dilutions of electroporated cells onto P. versutus agar containing 0.5% (vol/vol) methanol and the appropriate antibiotic (kanamycin or gentamicin). Optimization of the electroporation protocol was carried out with broad-host-range plasmid pJB3-Km1 (3).

Transposon mutagenesis of H. chloromethanicum.

Transposon mutants were generated by electroporation with plasposon pTnModRKm (8). Kanamycin-resistant colonies which appeared after 7 days of incubation were picked and grown in liquid cultures with 0.5% (vol/vol) methanol and kanamycin (50 μg ml⁻¹). Genomic DNA was extracted (37) from 50-ml cultures and analyzed for the presence of the transposon by PCR amplification of the kanamycin cassette and by Southern hybridization with the kanamycin cassette as a probe. The regions flanking the transposon insertions in CH₃Cl utilization mutants were cloned and sequenced as described previously (8). DNA from each CH₃Cl utilization mutant was extracted and digested with SphI, followed by ligation of different amounts of SphI-digested DNA (to allow the transposon with flanking regions to self-ligate and form a plasmid). Each ligation mixture was transformed into competent cells of E. coli CC118(λpir) by heat shock. E. coli CC118(λpir) carries a π gene required for replication from the RK6 origin of replication contained within the transposon (21), which enables circular molecules containing the transposon to replicate in this host. Following heat shock, the E. coli CC118(λpir) cells were plated onto kanamycin-supplemented agar and incubated for 16 h at 37°C. Kanamycin-resistant transformants containing circular molecules with the transposon and flanking regions were subjected to plasmid DNA preparations. The regions of DNA flanking the transposons were subsequently sequenced with primers designed to the outer regions of the transposon.

Selection procedure.

A colorimetric microtiter plate assay was developed to screen for CH₃Cl utilization-negative mutants of H. chloromethanicum. The assay is based on the addition of water-saturated phenol red (phenolsulfonphthalein) to a final concentration of 10% (vol/vol) in the growth medium. This changes color from red (pH 7.4) to yellow (pH 6.8) (5) during production of hydrochloric acid in the initial step of CH₃Cl utilization by H. chloromethanicum (Fig. 2).

FIG. 2.

Microtiter plate assay for selection of CH₃Cl-negative mutants of H. chloromethanicum. CH₃Cl-negative mutants of H. chloromethanicum were incubated in the presence of CH₃Cl in P. versutus medium containing 10% (vol/vol) water-saturated phenol red and kanamycin (50 μg/μl). For the incubation of wild-type H. chloromethanicum CM2, kanamycin was omitted. Following incubation, wells containing transposon mutants defective in CH₃Cl metabolism remained orange, whereas wells containing wild-type cells (WT) changed to a yellow color. Lanes 1 to 4 are four replicates of the same strain.

DNA manipulation.

Preparation of plasmid DNA, recombinant DNA work, and Southern analysis were performed with the methods of Sambrook and Russell (39). DNA was extracted from wild-type and mutant strains of H. chloromethanicum by the method of Oakley and Murrell (37). For Southern hybridization, DNA was transferred onto Hybond-N nylon membranes (Amersham) and fixed to the membrane with a UV Stratalinker (Stratagene). Probes were generated by PCR from the kanamycin cassette of plasmid pTnModRKm (8) and from the gentamicin cassette of plasmid p34S-Gm (8) and then purified from an agarose gel with a QIAquick gel extraction kit (Qiagen). DNA probes were labeled by random priming (10). The PCR-generated probe (50 ng) was labeled with 50 μCi of [α-³²P]dGTP, and unincorporated label was removed with the use of a MicroSpin column (Amersham Pharmacia Biotech), per the manufacturer's instructions. Prior to hybridization, probes were denatured in 0.4 M NaOH. Hybridization solution (0.5 M Na₂HPO₄, 0.5 M NaH₂PO₄ [pH 6.8] 7% [wt/vol] sodium dodecyl sulfate [SDS]) was used for 16 h at 60°C. Following hybridization, the membranes were washed twice in 2× SSC (17.3 g of NaCl and 8.8 g of trisodium citrate per liter, pH 7.0) at 60°C.

PCR.

PCR amplifications were performed in 50-μl (total volume) mixtures in 0.5-ml microcentrifuge tubes with a Hybaid Touchdown thermal cycling system. After an initial denaturation step at 94°C (5 min), 2.5 U of Taq polymerase (MBI Fermentas) was added. Amplification was carried out with 30 cycles of 94°C for 1 min, various annealing temperatures as required for 1 min, extension at 72°C for 1 min, and a final extension period at 72°C for 10 min.

DNA sequencing and analysis.

DNA sequencing was performed by cycle sequencing with a Dye Terminator kit (PE Applied Biosystems, Warrington, United Kingdom). DNA sequences were analyzed with a 373A automated sequencing system (PE Applied Biosystems). DNA sequences and derived amino acid sequences were analyzed with the DNAStar package. Similarity searches were performed with the gapped BLAST program (1) against public protein and gene databases (http://www.ncbi.nlm.nih.gov).

Isolation of total RNA.

RNA was extracted from 30 ml of early-exponential-phase (OD₅₄₀ = 0.3 to 0.5) cultures of H. chloromethanicum with the hot phenol method as previously described (12). Removal of DNA from the RNA preparations was done with 10 to 20 U of RQ1 DNase (Promega) according to the manufacturer's instructions. RNA quality was analyzed by running 2 μl of RNA preparation on a 1% (wt/vol) agarose gel in 2 μl of RNA loading buffer (1 ml formamide, 1 mg of xylene cyanol FF, 1 mg of bromophenol blue; 200 μl of 0.5 M EDTA, pH 8.0) and 8 μl of diethylpyrocarbonate (DEPC)-treated water. The presence of cmu-specific DNA fragments (RT regions RT1, RT2, RT3, RT4, and RT5 described in Fig. 6) was examined by PCR with primers specific for these regions.

FIG. 6.

RT-PCR products for the intergenic regions within the cmu cluster in H. chloromethanicum CM2. RNA was extracted from early-exponential cultures of H. chloromethanicum CM2 (wild type) grown on methanol, CH₃Cl or a mixture of methanol and CH₃Cl as described in the text. The black bands refer to the presence of the relevant RT-PCR product of intergenic regions RT1 to RT5 tested with the primers described in the text. The RT1 to RT5 regions amplified by RT-PCR are represented with black arrows.

RT-PCR.

The reverse transcription step was carried out with Expand reverse transcriptase (Roche) according to the manufacturer's instructions; 1 μg of RNA (DNA-free) was added to 50 pmol of reverse primer in 4.5 μl of DEPC-water and denatured at 65°C for 10 min, followed by cooling on ice for 2 min, and 2 μl each of 10 mM deoxynucleoside triphosphates, 2 μl of 100 mM dithiothreitol, 4 μl of 5× Expand buffer, 0.5 μl of DEPC-water, and 1 μl of Expand reverse transcriptase (40 U μl⁻¹) (Roche) were then added and incubated at 42°C for 1 h.

For RT-PCR, the following primers were used for the RT1 region (RT1-F, TCGTCGATCTTGGTGCAATG; RT1-R, CTATAAGTGCGGGCCAAACC), RT2 region (RT2-F, CAGCATCCTCGAGCATGCCGT; RT2-R, ACTCGATGTAGTCGCTACTAG), RT3 region (RT3-F, TGAGCAGTTCATCGAGAGTG RT; RT3-R, TCCATCGTACTGCAATCCAG), RT4 region (RT4-F, AGGTAGCATGACGTTAGGTC; RT4-R, CCGTTAAGTATCGGCATTCA), and RT5 region (RT5-F, TCACCTCTGGAAGCGATC; RT5-R, CCAATGTTGCCAGAACTA). PCRs were carried out as described above at an annealing temperature of 60°C (for RT regions 1 and 2) and 55°C (for RT regions 3, 4, and 5).

SDS-PAGE analysis.

Cells grown on CH₃Cl were harvested in the mid-exponential phase of growth (OD₅₄₀ = 0.5 to 0.7), washed once with minimal medium (P. versutus medium), resuspended in 0.1 M PIPES buffer (piperazine-N,N′-bis[2-ethanesulfonic acid], pH 7.0) and broken by three passages through a French pressure cell (American Instrument Company, Silver Spring, Md.) at 110 MPa (4°C). Cell debris was removed by centrifugation (35,000 × g, 25 min, 4°C). The resulting supernatant was used as a cell extract for SDS-polyacrylamide gel electrophoresis (PAGE) analysis. The protein content of the cell extract was determined with the Bio-Rad protein assay reagent according to the manufacturer's instructions. Protein samples (40 μg per lane) were analyzed by SDS-8% (wt/vol) PAGE (26) with an X-cell II Mini-Cell apparatus (Novex) and stained with Coomassie brilliant blue R250. Molecular masses were estimated with Dalton Mark VII-L molecular mass markers (Sigma).

Nucleotide sequence accession number.

The sequence of the extended cmu gene cluster of H. chloromethanicum has been deposited in the GenBank database under accession number AF281259.

RESULTS AND DISCUSSION

Optimization of the electroporation procedure.

The electroporation efficiency of H. chloromethanicum was optimized by altering the key parameters of the electroporation procedure (2, 28, 39, 43): the growth stage at which cells were harvested, the composition of the wash and electroporation buffers, and the parameters of the electrical pulse.

Glycerol (10%, vol/vol) as a wash buffer and electroporation buffer was previously used for the development of an electroporation procedure for Hyphomicrobium spp. (13, 53) and has been successfully applied in this study. Optimal electrical settings for the electrotransformation of H. chloromethanicum were previously determined to be 2.4 kV and 200 Ω at a capacitance of 25 μF (53). Maximum transformation efficiencies of 10⁶ transformants/μg of DNA were reproducibly obtained when cells were harvested at the early exponential phase of growth (OD₅₄₀ = 0.3). The transformation efficiency dropped dramatically to 10² transformants/μg of DNA when cells were harvested at the end of the exponential phase of growth.

This efficient electroporation procedure was further applied with the use of suicide transposon delivery vectors pUTmini-Tn5Km (21) and pTnModRKm (8). Significant transposition rates (10⁶ transformants/μg of DNA) were obtained with both vectors, which enabled electroporation to be used as a method of DNA transfer to create CH₃Cl utilization-negative mutants of H. chloromethanicum. This study has optimized the electroporation protocol for the efficient transformation of H. chloromethanicum, which may have widespread applicability for the genetic analysis of Hyphomicrobium spp.

Optimization of selection procedure for CH₃Cl-negative mutants.

Following electroporation, cells of H. chloromethanicum with transposon insertions in their genome were isolated by selection of the kanamycin resistance gene present on the transposon. Kanamycin-resistant cells in which insertional inactivation occurred in genes essential for CH₃Cl utilization were subsequently selected on the basis of their inability to grow on CH₃Cl. Due to the relatively slow growth of H. chloromethanicum on CH₃Cl, this selection procedure proved to be time-consuming (10 to 14 days) and inefficient.

A colorimetric microtiter plate assay, which relies on the production of HCl during degradation of CH₃Cl, was developed and used successfully to screen for CH₃Cl utilization-negative mutants of H. chloromethanicum (Fig. 2). The color change reaction (from red to yellow) of the assay is dependent on the production of hydrochloric acid by H. chloromethanicum during CH₃Cl metabolism. The microtiter plate system was selected in preference to solid medium as the color change in agar plates would affect the entire area rather than being restricted to a single colony. The microtiter plate was prepared by filling each well with 250 μl of P. versutus medium containing 10% (vol/vol) water-saturated phenol red and kanamycin (50 μg/μl). Kanamycin was omitted from wells prepared for the incubation of wild-type H. chloromethanicum CM2.

Following electroporation, kanamycin-resistant colonies were picked with sterile toothpicks onto master plates and into microtiter plates. Microtiter plates were incubated in sealed gas-tight jars gassed with 2% (vol/vol) CH₃Cl and incubated at 30°C. After incubation for 3 to 4 days, wells containing transposon mutants defective in CH₃Cl metabolism remained an orange color, whereas wells containing wild-type cells changed to a yellow color (Fig. 2). This selection procedure proved to be reliable, reproducible, time-efficient, and effective for selection of CH₃Cl utilization-negative mutants.

Identification and characterization of CH₃Cl utilization-negative H. chloromethanicum mutants.

From a total of 5,900 kanamycin-resistant transformants of H. chloromethanicum that were screened, 9 were unable to utilize CH₃Cl as their sole carbon and energy source. All nine CH₃Cl utilization-negative mutants were still able to grow with methanol, methylamine, formate, or acetate. The presence of the integrated transposon in the mutants was confirmed by probing SphI digests of total DNA from each mutant with the kanamycin resistance gene cassette (from pTn-ModRKm) as a probe (data not shown). Hybridization of the probe with DNA of all nine mutants resulted in a single hybridization fragment, indicating that the mutations were due to single-site insertions. The DNA fragments carrying a transposon insertion from each CH₃Cl-negative mutant were cloned and sequenced. Sequencing of DNA flanking the transposon was carried outwards, starting from the transposon through to the gene(s) that had been inactivated by mutagenesis. These sequencing data revealed the gene carrying the transposon insertion and provided a clear indication that the backbone of the suicide vector pTn-ModRKm did not integrate into the chromosome of any of the nine H. chloromethanicum mutants generated in these experiments.

Eight of the mutants carried a transposon insertion in cmuC, which encodes a putative methyltransferase, which resulted in the CH₃Cl-negative phenotypes of the mutants. The reason for the specific integration of the transposon into cmuC is not known, but the transposon targeted three different loci within the cmuC gene (Fig. 3). Analysis of the transposon insertion sites in all three groups of cmuC mutants did not reveal any specific flanking sequence repeats or palindromic sequences which might serve as a preferred target site for transposons (7, 22, 48). All eight cmuC mutants had lost the ability to grow on or dehalogenate CH₃Cl. Cell extracts of the cmuC mutants grown on methanol in the presence of CH₃Cl were further examined by SDS-PAGE alongside wild-type cells grown on CH₃Cl alone, a mixture of methanol and CH₃Cl, or methanol alone (Fig. 4).

FIG. 3.

Genes affected by mutagenesis in CH₃Cl utilization-negative mutants of H. chloromethanicum CM2. The positions of the transposon insertions in cmuC and hutI-metF mutants and the location of the gentamicin cassette in the cmuB⁻30 mutant are shown with solid black lines. Transposon insertions targeted three different locations within the cmuC gene, splitting cmuC mutants into three groups, as shown.

FIG. 4.

SDS-PAGE (8%) of CH₃Cl-induced 67-kDa protein from H. chloromethanicum CM2. Lanes: WT, wild type; MEOH, cells grown on methanol; CH₃Cl, cells grown on CH₃Cl; MEOH/CH₃Cl, cells grown on a mixture of methanol and CH₃Cl. hutI-metF⁻, cells of the hutI-metF mutant; cmuC⁻, cells of the cmuC mutant F30; cmuB⁻, cells of the cmuB⁻30 mutant. The arrow indicates the 67-kDa CH₃Cl-induced protein (CmuA). The amount of protein loaded per well was 40 μg.

SDS-PAGE analysis revealed that the cmuC mutants could not express the CH₃Cl-dependent 67-kDa polypeptide CmuA, whereas CH₃Cl-dependent expression of the 67-kDa polypeptide in wild-type cells grown on CH₃Cl or a mixture of methanol and CH₃Cl, but not in the methanol-grown wild type was clearly demonstrated (Fig. 4). This 67-kDa polypeptide corresponded to the predicted molecular mass of the derived CmuA polypeptide (67 kDa) and has been confirmed previously as CmuA by N-terminal sequence analysis (31). Absence of the CmuA polypeptide from cell extract of the methanol-grown wild type (Fig. 4) indicated CH₃Cl-dependent induction of the 67-kDa polypeptide (CmuA) in H. chloromethanicum. This is in accordance with previous experiments on H. chloromethanicum and M. chloromethanicum and Aminobacter sp. strain CC495, where CmuA expression is also CH₃Cl dependent (31, 45, 50). The absence of the 67-kDa polypeptide from cmuC mutants grown in the presence of CH₃Cl (Fig. 4) suggested that the mutation in the cmuC gene affected expression of the cmuA gene located downstream (3′) (Fig. 3). Such pleiotropic effects of mutations in cmuC on the cmuA gene located 3′ could be due to the organization of cmuA, cmuB, and cmuC, which probably constitute a single operon. The inability of the mutants to express CmuA, a methyltransferase catalyzing the dehalogenation step in CH₃Cl metabolism, would explain dechlorination-negative phenotypes of the cmuC mutants.

The cmuC gene was also found to be essential for CH₃Cl metabolism in M. chloromethanicum CM4, but the function of this gene remains unknown (50, 51). Similar studies have demonstrated that cmuC mutants of M. chloromethanicum were unable to grow with CH₃Cl, but did exhibit wild-type levels of both dehalogenase (CmuA) and methyltransferase II (CmuB) activity, indicating that the expression and functionality of both the CmuA and CmuB proteins were not affected by the mutation in cmuC (50). These differences could be explained by a different arrangement of the genes within the cmu cluster in H. chloromethanicum and within two unlinked cmu clusters in M. chloromethanicum (31, 32, 54). Although cmuB and cmuC are linked in M. chloromethanicum, cmuA is located on a separate transcriptional unit, and its expression is probably unaffected by mutation of cmuC (44, 50). In H. chloromethanicum, cmuB, cmuC, and cmuA are tightly linked on the chromosome and are possibly coregulated and coexpressed.

Sequencing of DNA flanking the transposon in mutant D20 indicated that the mutant carried a transposon insertion in the region containing the hutI and metF genes, which had not been defined in an earlier study (31) but which have been characterized in this study. Detailed analysis of the hutI-metF region indicated that the stop codon of the hutI gene overlapped the predicted start codon of the metF gene, suggesting that these two genes were likely to be transcriptionally coupled in H. chloromethanicum.

The precise location of the transposon was mapped to the 3′ end of the hutI gene, just four nucleotides upstream (5′) of the stop codon of hutI, which was also three nucleotides upstream of the start codon of the metF gene. Transposon integration into the hutI-metF region led to the dechlorination-negative phenotype for mutant D20, indicating that the mutation possibly affected expression of cmuA or of both cmuA and cmuB. SDS-PAGE analysis clearly revealed the inability of the hutI-metF mutant, like the cmuC mutant, to express the 67-kDa polypeptide (CmuA) (Fig. 4). Mutation in the hutI-metF region affected expression of the upstream (5′)-located cmuA and possibly all other genes located upstream (5′), which could possibly represent one transcriptional unit: cmuB, cmuC, cmuA, fmdB, paaE, hutI, and metF (Fig. 3). The exact mechanism of how a mutation in the hutI-metF region prevented expression of cmuA is not clear, but the data suggest that cmuB, cmuC, cmuA, fmdB, paaE, hutI, and metF in H. chloromethanicum are likely to be tightly coregulated and cotranscribed.

All previous attempts to detect metF in the genome of H. chloromethanicum by Southern hybridization with metF from M. chloromethanicum as a probe were unsuccessful (31). metF encodes a putative methylene-H₄-folate reductase, catalyzing the conversion of methyl-H₄-folate (formed from CH₃Cl during the dehalogenation step) into methylene-H₄-folate, a key intermediate of the CH₃Cl utilization pathway, which can subsequently be either oxidized to CO₂ or assimilated into cell material via the serine pathway (44, 50, 51, 54). Detection of metF in the genome of H. chloromethanicum provides evidence for the function of the H₄-folate-dependent CH₃Cl utilization pathway in this methylotroph.

Interestingly, cmu clusters from Aminobacter sp. strain IMB-1 and H. chloromethanicum (Fig. 3) exhibited an identical arrangement of cmuC, cmuA, fmdB, and paaE genes (32, 54). Moreover, in Aminobacter sp. strain IMB-1, paaE is 5′ of the hutI and metF genes, which indicated the possibility of the same gene order in H. chloromethanicum. PCR primers were designed to the end of paaE and to hutI and metF sequences in H. chloromethanicum obtained from the transposon mutagenesis experiment. Analysis of the PCR products produced confirmed that paaE was 5′ of hutI and metF on the cmu cluster of H. chloromethanicum and therefore extended the cluster by 2,546 bp (Fig. 3)

Sequence analysis of the extended CH₃Cl utilization cluster of H. chloromethanicum.

This study showed that paaE was 5′ of the previously undetected hutI and metF genes (Fig. 3). Complete sequencing of paaE in this study indicated that this gene in H. chloromethanicum (1,095 bp) encodes a 365-amino-acid polypeptide with significant identity (55%) to PaaE from Aminobacter sp. strain IMB-1. The paaE gene was not present in the two cmu clusters described for M. chloromethanicum (44). The derived PaaE polypeptide from H. chloromethanicum also showed 34, 30, and 28% identity to PaaE from Bradyrhizobium japonicum, E. coli, and Corynebacterium efficiens, respectively. PaaE from E. coli was reported to be the reductase which mediated electron transfer between NAD(P)H and the PaaACD oxygenase component during degradation of phenylacetic acid (11).

In H. chloromethanicum, hutI (1,311 bp) encodes a 437-amino-acid imidazolonepropionase with 35% identity to HutI from Aminobacter sp. strain IMB-1 (54). Unlike in H. chloromethanicum, the hutI gene located within the cmu cluster of Aminobacter sp. strain IMB-1 was smaller (1,019 bp) and encoded a 339-amino-acid polypeptide. Interestingly, the size of the imidazolonepropionase polypeptide encoded by hutI appeared to differ between Sinorhizobium meliloti (369 amino acids), Pseudomonas spp. (401 amino acids) and Burkholderia fungorum (431 amino acids). Although the link between HutI and metabolism of CH₃Cl has not yet been established, it is thought that in CH₃Cl utilizers, the imidazolonepropionase encoded by hutI may be involved in the formation of the imidazole ring found in the nucleotide loop of cobalamin (54).

The partial open reading frame metF (807 bp) in H. chloromethanicum (Fig. 3) encodes a methylene-H₄-folate reductase with 40% identity to MetF from Aminobacter sp. strain IMB-1 (54) and much lower identity (27%) to MetF from M. chloromethanicum (44, 50).

Marker exchange mutagenesis of cmuB.

To confirm the essential role of cmuB in H. chloromethanicum and to observe the effects of the cmuB knockout on the expression of genes located downstream of cmuB, a mutagenesis vector was constructed with a derivative of the suicide vector pK18mob (42). cmuB was previously demonstrated to encode a methyltransferase II (CmuB) involved in the dehalogenation of CH₃Cl in M. chloromethanicum (44, 45, 50). A 3.8-kb EcoRI-HindIII fragment containing the cmuB gene with flanking DNA was ligated into pK18mob. The cmuB gene was then disrupted by insertion of the gentamicin resistance cassette from p34S-Km (8) into a BamHI site which lies in the middle of cmuB (Fig. 5). This mutagenesis vector was transformed into H. chloromethanicum by electroporation with the optimized procedure described above.

FIG. 5.

Vector constructed for marker exchange mutagenesis of cmuB in H. chloromethanicum CM2. A 3.8-kb EcoRI-HindIII fragment containing the cmuB gene with flanking DNA was inserted into pK18mob. The cmuB gene was subsequently disrupted by insertion of the gentamicin resistance cassette (Gm^R) from p34S-Km into a BamHI site which lies within cmuB.

To inactivate cmuB from H. chloromethanicum, a double-crossover homologous recombination event between the constructed mutagenesis vector (Fig. 5) and the wild-type version of cmuB in the chromosome must occur. Of 100 gentamicin-resistant electrotransformants, 94 single-crossover mutants selected as gentamicin- and kanamycin-resistant colonies and six double-crossover mutants were isolated. Double-crossover mutants were identified as being gentamicin resistant and kanamycin sensitive, and strain cmuB⁻30 was chosen for further study. Analysis of strain cmuB⁻30 by PCR and Southern blotting (data not shown) indicated that a double-crossover event had occurred, resulting in loss of the plasmid backbone from the cell and insertion of the gentamicin cassette into cmuB.

Mutation of cmuB led to loss of the ability of H. chloromethanicum to grow on or to dehalogenate CH₃Cl. Cell extracts from strain cmuB⁻30 grown on a mixture of methanol and CH₃Cl were analyzed by SDS-PAGE (Fig. 4). This clearly indicated the absence of the CH₃Cl-induced 67-kDa polypeptide (CmuA). This confirmed that expression of the CmuA polypeptide encoded by cmuA was abolished by mutation of cmuB located 5′ of cmuA. Data obtained from both transposon and marker exchange mutagenesis studies led us to conclude that impaired expression of cmuA is likely to be due to polar effects of mutations in cmuB and cmuC, located upstream (5′) of cmuA within the operon cmuB, cmuC, cmuA, fmdB, paaE, hutI, and metF (cmuBCA-metF).

Transcriptional analysis of the cmu cluster of H. chloromethanicum.

RT-PCR analysis was carried out to assay for cmu-specific transcripts from total RNA extracted from H. chloromethanicum (wild type) grown on methanol alone, a mixture of methanol and CH₃Cl, or CH₃Cl alone. In addition, intergenic regions between cmuB and cmuC, cmuC and cmuA, cmuA and fmdB, fmdB and paaE, and hutI and metF were targeted in this RT-PCR study to investigate if genes within the cmu cluster of H. chloromethanicum were cotranscribed (Fig. 6). Positive controls with DNA from H. chloromethanicum as a template, PCR negative controls with water, and RT-PCR negatives to check for DNA contamination in RNA preparations were used in all cases. cDNA was synthesized from DNA-free RNA with an RT primer located within paaE (RT4 region) or metF (RT5 region) (Fig. 6).

Transcriptional analysis revealed RT-PCR products for all the intergenic regions tested within the cmu cluster from H. chloromethanicum (Fig. 6), indicating that all the genes within the cmu cluster (cmuB, cmuC, cmuA, fmdB, paaE, hutI, and metF) probably formed an operon (cmuBCA-metF) and were cotranscribed on a polycistronic mRNA. In addition, the transcript for the cmuBCA-metF operon was only present in cells grown on a mixture of methanol and CH₃Cl, or CH₃Cl alone and was absent from methanol-grown cells (Fig. 6). These RT-PCR data confirmed that expression of cmuB, cmuC, cmuA, fmdB, paaE, hutI, and metF was strictly CH₃Cl dependent. When H. chloromethanicum was grown on a mixture of methanol and CH₃Cl, methanol failed to repress expression of the cmuBCA-metF operon, indicating that transcription of the cmuBCA-metF operon is regulated solely by the presence or absence of CH₃Cl and is not repressed by an alternative carbon source. CH₃Cl-dependent gene expression of the cmuBCA-metF operon in H. chloromethanicum supports the SDS-PAGE data (Fig. 4) described previously, which showed that expression of the 67-kDa polypeptide (CmuA) was only detected in the presence of CH₃Cl (Fig. 4) (31).

Final conclusions.

This study provided improved methods for genetic analysis of Hyphomicrobium spp. Optimization of the electrotransformation protocol for H. chloromethanicum has considerable potential for the genetic analysis of other Hyphomicrobium spp. The successful application of electroporation to create stable Hyphomicrobium transposon and marker exchange mutants is reported for the first time. Development of the colorimetric microtiter plate assay in this study provided a facile screening mechanism for the identification of CH₃Cl utilization-negative mutants. Moreover, this colorimetric selection procedure could be widely applicable for the detection of CH₃Cl- or CH₃Br-negative mutants of other methylhalide-degrading bacteria. A suicide vector for specific gene (cmuB) inactivation has been constructed and used successfully to perform marker exchange mutagenesis in Hyphomicrobium spp. for the first time. This technique also provides a basis for future genetic manipulations with this genus.

This study provided strong evidence that H. chloromethanicum metabolizes CH₃Cl with the H₄-folate-dependent corrinoid-specific pathway that was initially detected in M. chloromethanicum (Fig. 1). Mutational analysis revealed that mutagenesis of cmuB, cmuC, or hutI had a detrimental effect on the functionality of the whole cmuBCA-metF operon and therefore on the whole CH₃Cl utilization pathway. All CH₃Cl-negative mutants of H. chloromethanicum were unable to grow on or oxidize or dehalogenate CH₃Cl or to express the CH₃Cl-dependent 67-kDa polypeptide (CmuA) (Fig. 4).

The mutagenesis data and transcriptional analysis of the cmu operon in H. chloromethanicum clearly implied that cmuB, cmuC, cmuA, fmdB, paaE, hutI, and metF genes were cotranscribed and coregulated. This provided a strong indication that, in dehalogenation-negative cmuB and cmuC mutants, defective expression of cmuA was likely to be due to polar effects of the inactivated cmuB or cmuC gene located upstream (5′) of cmuA within the cmuBCA-metF operon. The pleiotropic effect of the hutI-metF mutation on the expression of cmuA is not fully understood but provided further evidence that the cmuBCA-metF operon is coordinately expressed. It also suggested that disruption of any gene within the operon leads to loss of ability to grow on CH₃Cl. Expression of the cmuBCA-metF operon was strictly CH₃Cl dependent (Fig. 6). Such close transcriptional organization of the genes might suggest that transcription of the cmuBCA-metF operon operates from a single promoter, and we are currently attempting to locate this promoter.

Discovery of the metF and hutI genes in H. chloromethanicum within the cmuBCA-metF operon strongly suggested the functioning of the H₄-folate-dependent CH₃Cl oxidation pathway. It is likely that methyl-H₄-folate produced from CH₃Cl during the dehalogenation steps (catalyzed by CmuA and CmuB) is oxidized to formate via H₄-folate-linked intermediates (Fig. 1) and not via formaldehyde, as with other C₁ growth substrates used by the genus Hyphomicrobium (4, 14, 47).