Methane, the principal component of natural gas and biogas, is a cheap, abundant, and renewable energy and carbon source. However, methane is also the second most prevalent greenhouse gas (1). Therefore, technologies to efficiently convert methane to chemicals or fuels can bring new sustainable solutions to a number of industries with large environmental footprints. Methane-oxidizing bacteria (methanotrophs) are able to use methane as their sole source of carbon and energy and thus are promising systems for methane-based bioconversion (2, 3). The bioconversion of methane to industrial products (single-cell proteins, biopolymers, lipids, etc.) using aerobic methanotrophs has been studied for approximately 50 years but has had little enduring success (4). Current biological engineering and systems biology approaches provide new opportunities for metabolic engineering of methanotrophs. However, to be an industrial workhorse like Escherichia coli, an industrial methanotroph needs to have a high growth rate, and efficient genetic tools for its manipulation and a fundamental knowledge base are needed.
Many methanotrophs have been isolated and characterized since the classic study of Whittenbury et al. (5). Aerobic methanotrophs are found in two major groups, the gammaproteobacterial methanotrophs (type I) and the alphaproteobacterial methanotrophs (type II) (2). Most studies and biotechnological efforts have focused on well-characterized species, such as Methylococcus capsulatus Bath, Methylosinus trichosporium OB3b, and Methylocystisparvus OBBP (6). In recent years, a variety of new strains have been isolated, including thermophilic, psychrophilic, acidophilic, alkaliphilic, and halophilic methanotrophs, which have expanded the physiological range of aerobic methanotrophs (7, 8). Moreover, some isolates show robust growth and promise for use in biotechnological applications. Type I methanotrophs are particularly well suited for use in industrial processes, as they utilize the highly efficient ribulose monophosphate pathway for formaldehyde assimilation into biomass (2, 6). A particularly promising type I methanotroph for industrial use is Methylomicrobium buryatense 5G, a haloalkaliphilic strain that was isolated directly on natural gas from a soda lake in the Transbaikal region of Russia, a harsh environment with fluctuating temperatures, salinities, and pH levels (7). The pH optimum of this strain is 9.5, and the optimum salt concentration is 0.75%. M. buryatense strain 5GB1, a lab-adapted variant of M. buryatense 5G, grows well in pure culture on natural gas, methane, and methanol (9). The genomes of M. buryatense 5G and 5GB1 were recently sequenced and annotated (10). All of these parameters make this strain well suited for development as an industrial system for the bioconversion of methane.
Historically, the slow pace of genetic manipulation has been one of the limiting factors in analyzing the gene functions of methanotrophic bacteria. At present, conjugation is a method commonly used for the transformation of methanotrophs (6). Recently, to metabolically engineer M. buryatense 5GB1, Puri et al. developed a number of genetic tools for use with the strain, including a sucrose counterselection system and a small replicable vector, and these were found to be sufficient for genetic manipulation (9). However, as these tools are based on conjugation and vector construction, a number of steps are necessary to carry out genetic manipulation, making current techniques time-consuming. To accelerate the use of modern metabolic engineering techniques in M. buryatense 5GB1 and other type I methanotrophs, more streamlined and versatile approaches are needed. Genetic manipulation via electroporation requires fewer steps than conjugation and creates the ability to transfer linear DNA fragments, which results in double-crossover deletion and insertion events and removes the need for the counterselection of E. coli postconjugation. In addition, it allows the direct transfer of genes that might be toxic in alternate hosts, such as E. coli, and avoids the alteration of plasmids occurring during conjugation (9). Electroporation protocols have been developed in Methylocystis sp. strain SC2 and Methylocella silvestris BL2, both of which are type II methanotrophs (11, 12). However, no electroporation method has been reported for type I methanotrophs, and previous attempts to demonstrate electroporation in M. buryatense were unsuccessful (9).
In this work, we report simple and rapid electroporation-based genetic manipulation tools for multiple type I methanotrophs, including electroporation of DNA fragments that were assembled by PCR. We were able to obtain a high efficiency of DNA transfer via electroporation in M. buryatense strain 5GB1C, Methylomonas sp. strain LW13, and Methylobactertundripaludum strain 21/22, type I methanotrophs from three different genera with different physiological traits (9, 13), demonstrating the broad applicability of the techniques developed here. In M. buryatense, we also demonstrate marker removal after electroporation through sacB counterselection or FLP-FRT (FLP recombination target) site-specific recombination. These tools were validated by performing gene knockouts, long fragment deletion, and integration of a foreign fragment into the chromosome. These genetic tools will enable the rapid genetic manipulation and metabolic engineering of type I methanotrophs, accelerating the study of methanotrophs and their development for use in industrial processes.
MATERIALS AND METHODS
Bacterial strains, plasmids, oligonucleotides, and DNA manipulation techniques.
The bacterial strains and plasmids used in this study are listed in Table 1. All M. buryatense strains were derived from M. buryatense strain 5G (7). Strain 5GB1, a variant of M. buryatense 5G, shows a higher growth rate than its parent strain, and its sequence has several differences, identified by genome sequencing, from the published draft genome sequence of strain 5G (9). Strain 5GB1C, a variant of strain 5GB1 intentionally cured of its native plasmid, is capable of being conjugated with small IncP-based plasmids (9). The oligonucleotides (see Table S1 in the supplemental material) were synthesized by Integrated DNA Technologies (IDT; USA), and DNA sequencing was performed by Genewiz (USA). All plasmids were constructed by use of Gibson assembly (14). Phusion DNA high-fidelity polymerase was purchased from New England Biolabs (NEB; USA). A QIAquick PCR purification kit and a QIAquick gel extraction kit were supplied by Qiagen (Germany). Gene identities are from GenBank.
Culture, growth conditions, and phenotypic characterization.
All strains were cultured in an atmosphere of 25% methane in air at 30°C (M. buryatense 5GB1C) or 20°C (Methylomonas sp. LW13 and M. tundripaludum 21/22). The plates were incubated in sealed jars (Oxoid Limited, Hampshire, United Kingdom), while liquid cultures were grown in 250-ml glass serum bottles (Kimble Chase, Vineland, NJ, USA) sealed with rubber stoppers and aluminum seals (Wheaton, Millville, NJ, USA). Milli-Q H2O (Barnstead) was used in this study. M. buryatense cultures were grown in NMS2 medium (9), and Methylomonas and Methylobacter cultures were grown in NMS medium (15). NMS medium and NMS2 medium were supplemented with 1.5% Bacto agar for plate growth. The following antibiotics were used for the selection of colonies of both methanotrophs and E. coli containing the correct genetic manipulations: kanamycin (Km), 50 μg/ml; rifamycin (Rm), 50 μg/ml; hygromycin (Hm), 100 μg/ml; and zeocin (Zeo), 30 μg/ml. To detect the activity of soluble methane monooxygenase (sMMO), M. buryatense strains were first grown in NMS2 medium without copper at 30°C for 20 h and then subjected to the colorimetric plate assay developed by Graham et al. (16).
Electroporation protocol for type I methanotrophs.
For cells grown on agar plates, one loopful of cells was spread across an entire NMS or NMS2 agar plate and grown under methane overnight (M. buryatense 5GB1C and Methylomonas sp. LW13) or for 2 days (M. tundripaludum 21/22). For cells grown in liquid, a 50-ml liquid culture was grown to stationary phase as described above. Cells were harvested from the liquid culture by centrifugation at 5,000 rpm and 4°C for 10 min. Cells were harvested from the plates by scooping the entire biomass from a plate. Cells were resuspended in 50 ml room temperature electroporation solution (H2O, 9.3% sucrose, 10% glycerol, or 30% polyethylene glycol [PEG] 6000) and harvested by centrifugation at 5,000 rpm and 4°C for 10 min. The pellet was washed in 10 ml room temperature electroporation solution, transferred to a 15-ml conical tube, and centrifuged again at 5,000 rpm and 4°C for 10 min. The resulting pellet was resuspended in 100 to 150 μl electroporation solution and placed on ice. Fifty microliters of the cell suspension was mixed gently with DNA (usually less than 3 μl), and the mixture was transferred to an ice-cold 1-mm-gap cuvette (Bio-Rad). Electroporation was performed using a Gene Pulser II system (Bio-Rad) set at 25 μF and 200 Ω. Immediately following the electrical discharge, 1 ml of medium (room temperature) was added to the cells. The resuspended cells were then transferred into 10 ml medium in 250-ml serum bottles, which were crimp sealed, and then incubated with either 50 ml methane or 0.02% methanol. After incubation at 30°C (M. buryatense 5GB1C) or 20°C (Methylomonas sp. strain LW13 and M. tundripaludum 21/22) for 4 h or overnight with shaking, the cells were centrifuged at 5,000 rpm for 10 min at room temperature, resuspended in 1 ml medium, and spread onto selective plates.
Fusion of multiple DNA fragments by PCR.
Fusion of multiple DNA fragments by PCR was performed as described by Shevchuk et al. (17). In brief, it was carried out as follows. Overlaps of 30 to 40 nucleotides were introduced between each of two fragments through the use of primers. The fragments were amplified and gel purified. The reaction mixture in step A was 9.5 μl water, 5 μl Phusion buffer (5×) (NEB, USA), 2 μl deoxynucleoside triphosphate (dNTP) mix (2.5 mM each), 8 μl gel-purified fragments (about 50 ng each), and 0.5 μl Phusion DNA polymerase; the cycling parameters were an initial denaturation at 98°C for 2 min and subsequent steps of 98°C for 10 s, annealing at 55°C for 20 s, and extension at 72°C for 2 min for 16 cycles total. The reaction mixture in step B was 32 μl water, 10 μl Phusion buffer, 4 μl dNTP mix, 2 μl forward and reverse primers (10 mM) specific for the expected fragment, 1 μl of the unpurified PCR product from step A, and 1 μl Phusion DNA polymerase; the cycling parameters were an initial denaturation at 98°C 2 for min and subsequent steps of 98°C for 10 s, annealing at 60°C for 10 s, and extension at 72°C for 2 min for 32 cycles total. The resulting product was purified by use of a QIAquick PCR purification kit (Qiagen), and the purified product was directly electroporated into the strains.
Construction of ZS cassette.
The zeocin resistance marker (Zeor) was generated through two-step PCR amplification from plasmid p7Z6 (18) using primer pairs z51/z3 and z52/z3. The fragment containing the tac promoter (Ptac) and the Shine-Dalgarno (SD) sequence of lacZ from pCM66 (19) was introduced to the 5′ end of the zeocin resistance gene Streptoalloteichus hindustanus (Sh) ble with the primers z51 and z52. To assemble the Zeor-sacB (ZS) cassette, the sacB gene together with its putative SD sequence was fused to the 3′ end of Zeor with primers z52, z3, s5, and s3.
Introduction of Ppmo-xylE into the 5GB1C chromosome.
The pFC25 vector was constructed by use of Gibson assembly using a hygromycin-resistant version of the replicable vector pCM158 as a backbone (20). The hygromycin resistance marker (Hmr) was amplified by PCR from plasmid pTEC27 (Addgene accession number 30182) (21), the flippase gene was amplified by PCR from plasmid pE-FLP (Addgene accession number 45978) (22), and Ptac was amplified from pAWP89 (9). To create the insertion PCR product containing the pmo promoter fused to xylE, two PCR products were generated for the flanking integration sites and correspond to the intergenic region and coding sequences of the genes METBUDRAFT_2794 and METBUDRAFT_2795. PCR products corresponding to the pmo promoter, xylE, and Zeor flanked by FRT sites were inserted between the flanking regions via the fusion PCR method. The xylE gene was amplified from pCM130 (19). FRT sites were appended to Zeor by amplifying Zeor with primers containing the FRT sites.
Deletion of R-M systems and in vitro methylation of plasmid DNA.
Ten restriction-modification (R-M) system-associated restriction endonuclease genes (see Table S2 in the supplemental material) were predicted to exist in the genome of M. buryatense 5G through analysis of the restriction enzyme database REBASE (23) and bioinformatics analysis. Among these restriction endonuclease genes, METBUDRAFT_0633, METBUDRAFT_3180, and METBUDRAFT_4274 (the gene locus tags in the Integrated Microbial Genomes system [img.jgi.doe.gov/]) were deleted through unmarked allelic exchange using plasmid pCM433kanT (9), and others were deleted by electroporating the fusion PCR product containing the kanamycin resistance marker and two flanking homology regions of the target gene.
The methyltransferase gene METBUDRAFT_0046 was ligated to pET-29b(+) between the NdeI and XhoI sites, generating pET29-M. This vector contains a His tag sequence that is added to the C terminus of the expressed protein. E. coli BL21-AI harboring pET29-M was grown at 37°C in Luria-Bertani broth supplemented with Km to an optical density at 600 nm of ∼0.5 and induced with 0.1% arabinose and 0.5 mM IPTG (isopropyl-β-d-thiogalactopyranoside) at 30°C for 4 h. His-tagged (C-terminal) protein was purified using Talon metal affinity resin (Clontech) according to the supplier's instructions. For in vitro methylation, 10 mg of purified protein was incubated with 50 mM Tris-HCl (pH 8.5), 50 mM NaCl, 80 mM S-adenosylmethionine, 10 mM dithiothreitol, 10 mM EDTA, and 10 μg plasmid DNA substrate in a 200-μl reaction mixture for 2 h, 4 h, or 8 h at 30°C. Methylated plasmid DNA was purified and concentrated by phenol-chloroform extraction and ethanol precipitation.
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