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Journal of Bacteriology, March 2008, p. 1928-1936, Vol. 190, No. 6
0021-9193/08/$08.00+0     doi:10.1128/JB.01424-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Mutagenesis of the C1 Oxidation Pathway in Methanosarcina barkeri: New Insights into the Mtr/Mer Bypass Pathway

Paula V. Welander and William W. Metcalf^*

Department of Microbiology, University of Illinois at Urbana-Champaign, B103 Chemical and Life Science Laboratory, 601 South Goodwin Avenue, Urbana, Illinois 61801

Received 3 September 2007/ Accepted 23 December 2007

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A series of Methanosarcina barkeri mutants lacking the genes encoding the enzymes involved in the C1 oxidation/reduction pathway were constructed. Mutants lacking the methyl-tetrahydromethanopterin (H₄MPT):coenzyme M (CoM) methyltransferase-encoding operon (mtr), the methylene-H₄MPT reductase-encoding gene (mer), the methylene-H₄MPT dehydrogenase-encoding gene (mtd), and the formyl-methanofuran:H₄MPT formyl-transferase-encoding gene (ftr) all failed to grow using either methanol or H₂/CO₂ as a growth substrate, indicating that there is an absolute requirement for the C1 oxidation/reduction pathway for hydrogenotrophic and methylotrophic methanogenesis. The mutants also failed to grow on acetate, and we suggest that this was due to an inability to generate the reducing equivalents needed for biosynthetic reactions. Despite their lack of growth on methanol, the mtr and mer mutants were capable of producing methane from this substrate, whereas the mtd and ftr mutants were not. Thus, there is an Mtr/Mer bypass pathway that allows oxidation of methanol to the level of methylene-H₄MPT in M. barkeri. The data further suggested that formaldehyde may be an intermediate in this bypass; however, no methanol dehydrogenase activity was found in mtr cell extracts, nor was there an obligate role for the formaldehyde-activating enzyme (Fae), which has been shown to catalyze the condensation of formaldehyde and H₄MPT in vitro. Both the mer and mtr mutants were able to grow on a combination of methanol plus acetate, but they did so by metabolic pathways that are clearly distinct from each other and from previously characterized methanogenic pathways.

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Biologically mediated methanogenesis is carried out by a diverse group of anaerobic organisms, known as methanoarchaea, that obtain all of their energy for growth by converting a limited number of substrates to methane (CH₄) (5). These organisms are found in a variety of environments throughout the world, including both man-made and natural anaerobic habitats in which there are limited amounts of light, sulfate, and nitrate but which contain complex organic compounds (4). Despite the enormous phylogenetic diversity among the methanogens, the metabolism of these organisms is extremely specialized, and all known species are obligate methanogens. Four discrete methanogenic pathways are known: (i) the CO₂ reduction or hydrogenotrophic pathway involves the reduction of CO₂ to CH₄ with hydrogen gas (H₂) as the reductant; (ii) the methylotrophic pathway involves the disproportionation of methylated compounds, such as methanol and methylamines, to CO₂ and CH₄; (iii) the methyl reduction pathway results in the direct reduction of methanol to CH₄ with H₂ as the reductant; and (iv) the acetoclastic pathway involves the dismutation of acetate to CO₂ and CH₄ (7). A key series of reactions in both the methylotrophic and hydrogenotrophic pathways involves the reversible oxidation/reduction of coenzyme-bound one-carbon units (the C1 oxidation/reduction pathway). Extensive in vitro biochemical studies over the years have characterized the unique proteins involved in this pathway, leading to elucidation of the pathway shown in Fig. 1 (4, 7, 23). However, in vivo studies of the pathway were not possible until the relatively recent development of tools for facile genetic manipulation of Methanosarcina species (21).

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FIG. 1. Methylotrophic methanogenesis in M. barkeri Fusaro. Methylated compounds, such as methanol, can be disproportionated by M. barkeri to CO2 and CH4. In this pathway, one molecule of methanol is oxidized to CO2 to provide the reducing equivalents necessary to reduce three additional molecules of methanol to CH4. In this study, single-deletion mutants were constructed for the genes encoding four of the proteins required in the C1 oxidation branch of the methylotrophic pathway (Ftr, Mtd, Mer, and Mtr). Abbreviations: CHO-MF, formyl-methanofuran; CHO-H4MPT, formyl-H4MPT; CHH4MPT, methenyl-H4MPT; CH2H4MPT, methylene-H4MPT; CH3-H4MPT, methyl-H4MPT; CH3-CoM, methyl-CoM; CoM-CoB, mixed disulfide of CoM and CoB; MePh, oxidized methanophenazine, MePhH2, reduced methanophenazine; F420, F420; F420H2, reduced F420; Fd(ox), oxidized ferredoxin; Fd(red), reduced ferredoxin; Ftr, formyl-methanofuran:H4MPT formyltransferase; Mch, methenyl-H4MPT cyclohydrolase; Mtd, methylene-H4MPT dehydrogenase; Mer, methylene-H4MPT reductase; Mtr, methyl-H4MPT:CoM methyltransferase.

Genetic studies of the C1 oxidation/reduction pathway in Methanosarcina barkeri have shown that some of the steps have multiple roles in the physiology of the cell. For example, studies of mutants lacking the enzyme methenyl-tetrahydromethanopterin (methenyl-H₄MPT) cyclohydrolase (Mch), which catalyzes the conversion of methenyl-H₄MPT to formyl-H₄MPT, indicate that the C1 pathway is required both for energy production and for biosynthesis (11). Genetic studies have also indicated that there are as-yet-uncharacterized biochemical pathways that may allow some steps of the C1 oxidation pathway to be bypassed.

The energy-consuming methyl transfer from methyl coenzyme M (methyl-CoM) to H₄MPT is thought to be a critical step in the C1 oxidation/reduction pathway. This reaction is catalyzed by the N⁵-methyl-H₄MPT:CoM methyltransferase (Mtr), an eight-subunit membrane-bound enzyme encoded by the mtrECDBAFGH operon (10). We have demonstrated that Mtr is dispensable in M. barkeri under a variety of conditions but is required for growth via the hydrogenotrophic, aceticlastic, and methylotrophic pathways (26). Interestingly, although a mtr deletion mutant is unable to utilize methanol as a growth substrate, mtr cells are still able to oxidize methanol to CO₂, clearly demonstrating that the Mtr-catalyzed reaction must be bypassed in M. barkeri. Mutants lacking methenyl-H₄MPT cyclohydrolase (Mch) are unable to produce CH₄ from methanol in cell suspensions (11). Because the mtr mutants are capable of methanol oxidation and the mch mutants are not, the bypass must join the standard pathway somewhere between methenyl-H₄MPT and methyl-H₄MPT.

Finally, analysis of C1 oxidation/reduction pathway mutants also has indicated that there may be previously undiscovered methanogenic pathways in Methanosarcina. Although unable to grow using either methanol or acetate, mtr mutants are capable of growing on a combination of these two substrates. Labeling studies showed that growth on this combination of substrates occurred via a new methanogenic pathway in which acetate oxidation was used to drive methanogenic reduction of methanol. Because Mtr activity is sodium dependent, it seems possible that this pathway may be physiologically relevant in low-sodium environments (26).

These unexpected observations clearly stress the need for further in vivo analysis of methanogenesis. Here we report construction and characterization of additional mutants with mutations in the C1 oxidation pathway. The results of our studies clearly establish the role of the steps in the C1 oxidation pathway during growth of M. barkeri via the hydrogenotrophic, methylotrophic, and aceticlastic pathways and shed new light on the physiological role of the Mtr bypass pathway.

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Bacterial strains, media, and growth conditions. Escherichia coli strains were grown under standard conditions (25). M. barkeri strains were grown with single-cell morphology (22) at 37°C under strictly anaerobic conditions in high-salt (HS) medium (18). HS media were supplemented when necessary with methanol (125 mM) or acetate (80 mM alone or 40 mM in combination with methanol) under either an N₂-CO₂ (80:20; 40 kPa over ambient pressure) or H₂-CO₂ (80:20; 150 kPa over ambient pressure) headspace. Bacteria were grown on HS media solidified with 1.5% agar as described by Boccazzi et al. (2). All plating manipulations were carried out under strictly anaerobic conditions in an anaerobic glove box. Plates containing solid media were incubated in an intrachamber anaerobic incubator as described previously (19). Puromycin (Calbiochem, San Diego, CA) was added from sterile anaerobic stocks at a final concentration of 2 µg/ml for the selection of M. barkeri strains carrying the puromycin transacetylase gene (pac). The purine analog 8-aza-2,6-diaminopurine (8-ADP) (Sigma, St. Louis, MO) was added from sterile, anaerobic stock solutions at a final concentration of 20 µg/ml for selection against the hpt gene.

DNA methods, plasmid construction, and transformation. Standard methods were used throughout this study for isolation of plasmid DNA from E. coli (1). All plasmids and primers used in this study are described in Table 1 and Table 2. Genomic DNA from M. barkeri strains was isolated as described previously (20). DNA hybridization was performed using the DIG system (Roche, Mannheim, Germany) with magnagraph nylon transfer membranes obtained from Osmonic (Westborough, MA). DNA sequences of all cloning intermediates were confirmed at the W. M. Keck Center for Comparative and Functional Genomics, University of Illinois. E. coli strains were transformed by electroporation using an E. coli Gene Pulser (Bio-Rad, Hercules, CA) as recommended by the supplier. Liposome-mediated transformation was used for Methanosarcina as described previously (2, 17).

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TABLE 1. Plasmids used in this study

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TABLE 2. Primers used in this study

Construction of M. barkeri deletion strains. M. barkeri deletion strains (Table 3) used in this study were derivatives of WWM85 or WWM86 and were constructed by homologous recombination-mediated gene replacement as described previously (21). Briefly, 2 kb of the region 5' of the gene of interest and 2 kb of the region 3' of the gene of interest were PCR amplified utilizing KOD DNA polymerase (Novagen, Madison, WI) and M. barkeri Fusaro (= DSM 804) genomic DNA as the template. Each region was cloned into pJK301 (21) on either side of a pac-hpt cassette that was flanked by two 34-bp Frt sites in the same orientation. Two micrograms of the deletion plasmid was linearized by digestion with NotI and transformed into 8-ADP-resistant M. barkeri strain WWM85. Transformants were selected on HS agar containing methanol and puromycin under an H₂-CO₂ gas phase. Puromycin-resistant strains were single colony purified and screened by PCR for replacement of the wild-type locus with the pac-hpt cassette. The pac-hpt cassette was subsequently removed by transforming the resulting deletion strain with 2 µg of the nonreplicating plasmid pMR55 (21), which contains the Flp recombinase gene under control of the mcrB promoter from M. barkeri Fusaro, and plating the preparation on HS agar with methanol and 8-ADP under an H₂-CO₂ gas phase. 8-ADP-resistant clones were screened by PCR, and removal of the pac-hpt cassette was verified by Southern blotting.

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TABLE 3. M. barkeri strains used in this study

Cell suspension experiments. M. barkeri strains were grown on methanol under an H₂-CO₂ gas phase to late exponential phase (optical density at 600 nm [OD₆₀₀], 0.6 to 0.7). Cells were harvested by centrifugation at 5,000 x g for 15 min at 4°C. Cells were washed twice with anaerobic HS PIPES buffer [50 mM piperazine-N,N'-bis(2-ethansulfonic acid) (pH 6.8), 400 mM NaCl, 13 mM KCl, 54 mM MgCl₂, 2 mM CaCl₂, 2.8 mM cysteine, 0.4 mM Na₂S], resuspended in the same buffer to a final concentration of 10⁹ cells per ml, and kept on ice. All assay mixtures contained 2 ml of the cell suspension (2 x 10⁹ cells), and the assays were carried out under strictly anaerobic conditions in 25-ml Balch tubes sealed with butyl rubber stoppers. Puromycin (20 µg/ml) was added to prevent protein synthesis, and each assay mixture contained either 250 mM methanol or 250 mM acetate or both. The gas phase was exchanged five times with N₂ or H₂ and pressurized to 150 kPa over the ambient pressure. Cells were kept on ice until they were used, and assays were started by transferring tubes to 37°C and incubating them on a roller drum (New Brunswick Scientific).

For rate determination, gas phase samples (0.1 ml) were withdrawn with a gas-tight sample lock Hamilton syringe every 10 min, and CH₄ was assayed by gas chromatography (GC) at 225°C with a Hewlett Packard gas chromatograph (5890 series II) equipped with a flame ionization detector. The column used was a stainless steel column filled with 80/120 Carbopack B/3% SP-1500 (Supelco, Bellefonte, PA), and helium was used as the carrier gas. To determine total CH₄ and CO₂ production, assay mixtures were incubated at 37°C for 48 h, and then 0.1-ml gas phase samples were withdrawn. These samples were analyzed by GC at 225°C using a Hewlett Packard gas chromatograph (5890 series II) equipped with a thermal conductivity detector. A stainless steel 60/80 Carboxen-1000 column (Supelco) with helium as the carrier gas was used for these determinations. The total cell protein content was determined using the Coomassie blue protein assay reagent (Pierce, Rockford, IL) after an aliquot of the cells was lysed by sonication for 10 s.

Labeled cell suspension experiments. Cell suspension assays were performed as described above, except that each assay mixture contained 250 mM [¹³C]methanol (Isotec, Miamisburg, OH). Gas phase samples (0.1 ml) were removed after 48 h of incubation on roller drum at 37°C, and the ¹³CH₄ and ¹³CO₂ contents were determined by gas GC-mass spectrometry (MS). This experiment was done at 50°C using an HP6890N GC system equipped with an HP5973 mass selective detector, a flame ionization detector, and a thermal conductivity detector. A carbon plot capillary column (length, 30 m; inside diameter, 0.32 mm; Agilent Technologies, Colorado Springs, CO) was used with helium at flow rate of 2.5 ml/min as the carrier gas.

To determine how mer cells metabolized methanol plus acetate, cell suspension assays were performed as described above, with the following changes. mer cells were grown for 3 months in 250 ml of HS medium containing 250 mM methanol plus 160 mM acetate. Cell suspensions were prepared in triplicate, and one of the following combinations was added to each suspension: 250 mM [¹³C]methanol plus unlabeled 250 mM acetate, unlabeled 250 mM methanol plus 250 mM [1-¹³C]acetate, or unlabeled 250 mM methanol plus 250 mM [2-¹³C]acetate. Cell suspensions were incubated at 37°C for 5 days, and assays to determine the ¹³CH₄ and ¹³CO₂ contents were performed by GC-MS as described above.

Methyltransferase assays. Wild-type and fae strains were grown on methanol to late log phase (OD₆₀₀, 0.6 to 0.7), and cell extracts were then prepared anaerobically as previously described (26). Methyltransferase activity in these extracts was determined by measuring the formation of methyl-CoM from formaldehyde, H₂, and CoM as previously described (26).

Methanol dehydrogenase assays. mtr cells were grown on methanol plus acetate to late log phase (OD₆₀₀, 0.6 to 0.7), and cell extracts were then prepared anaerobically as described previously (26). Methanol dehydrogenase activity was assayed in both the reductive and oxidative directions. For the oxidative direction, each assay mixture contained 100 mM methanol, crude extract (1 mg of protein), 5 mM 2-bromoethane sulfonic acid, and one of the following electron acceptors: 20 µM factor 420 (F₄₂₀), 100 µM flavin adenine dinucleotide (FAD), 100 µM flavin mononucleotide (FMN), 100 µM pyrroloquinoline quinine (PQQ), 5 mM methyl viologen (MV), 5 mM benzyl viologen (BV), 100 µM 2,6-dichlorophenol indophenol (DCPIP), 200 µM metronidazole (MND), 1 mM phenazine methosulfate (PMS), or 20 µM cytochrome c (equine heart). Plumbagin (5-hydroxy-2-methyl-1,4-naphthoquinone) (100 µM) was also used as an electron acceptor, but the assay mixture also contained 20 µM cytochrome c to monitor the reduction of the quinone spectophotometrically. Electron acceptors were purchased from Sigma (St. Louis, MO). F₄₂₀ was a gift from L. Daniels.

For the reductive direction, each assay mixture contained either 10 or 1 mM formaldehyde, prepared from paraformaldehyde as described previously (26), crude extract (1 mg of protein), 5 mM 2-bromoethane sulfonic acid, and one of the following electron donors: 200 µM NADH, 200 µM NADPH, 20 µM reduced F₄₂₀, 100 µM reduced PQQ, 100 µM reduced FAD, 100 µM reduced FMN, 100 µM reduced DCPIP, 5 mM reduced BV, 5 mM reduced MV, 20 µM reduced MND, or 1 mM reduced PMS. NADH and NADPH were purchased from Sigma (St. Louis, MO). F₄₂₀ was reduced in the anaerobic glove box by adding 0.08 g of borohydride polymer (2.5 mmol/g; Sigma, St. Louis, MO) to 600 µl of 100 µM F₄₂₀ and incubating the preparation at room temperature for 30 min. After the 30-min incubation, the solution of F₄₂₀ appeared to be colorless and had no absorbance at 400 nm, indicating that the F₄₂₀ had been reduced. The borohydride was removed by centrifugation for 1 min at the maximum speed. Reduced F₄₂₀ was transferred to a new tube, and 0.2 ml was used in each assay mixture. All other electron carriers were reduced by preparing a 10x stock solution in 50 mM anaerobic morpholinepropanesulfonic acid (MOPS) and adding a few crystals of palladium black (Sigma, St. Louis, MO). The tubes were sealed, and the gas was exchanged five times with 100% N₂ to remove any remaining oxygen. A gas mixture containing 10% H₂ and 90% N₂ was then added by exchanging it five times and pressurized to 150 kPa over ambient pressure. The stock solutions were then incubated at room temperature for 1 h with shaking to allow reduction of the cofactor. The reduced cofactor was then anaerobically transferred to a microcentrifuge tube and centrifuged for 1 min at the maximum speed to remove the palladium black; 0.1 ml of the supernatant containing the reduced cofactor was then used in each assay.

Each methanol dehydrogenase assay was set up anaerobically using a 1-ml (final volume) mixture with 50 mM MOPS (pH 7.0). The cuvette was sealed with a stopper, and the gas phase was exchanged five times with 100% N₂. Each assay was begun by anaerobically adding either methanol or formaldehyde from an anaerobic stock solution using a Hamilton syringe, and the activity was monitored by following the reduction or oxidation of each electron acceptor or donor at 37°C at the following wavelengths: F₄₂₀, 400 nm ( = 25.7 mM^–1 cm^–1); NAD, 340 nm ( = 6.3 mM^–1 cm^–1); NADP, 340 nm ( = 6.3 mM^–1 cm^–1); FAD, 450 nm ( = 11.3 mM^–1 cm^–1); FMN, 450 nm ( = 11.3 mM^–1 cm^–1); PQQ, 302 nm ( = 25.1 mM^–1 cm^–1); MV, 600 nm ( = 8.3 mM^–1 cm^–1); BV, 500 nm ( = 7.7 mM^–1 cm^–1); DCPIP, 600 nm ( = 16 mM^–1 cm^–1); MND, 320 nm ( = 9.2 mM^–1 cm^–1); PMS, 387 nm ( = 26.3 mM^–1 cm^–1); and cytochrome c, 550 nm ( = 29.5 mM^–1 cm^–1).

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Construction of M. barkeri methyl oxidation deletion mutants. Unmarked deletion mutants with mutations in the genes encoding methyl-H₄MPT:CoM methyltransferase (mtr), methylene-H₄MPT reductase (mer), methylene-H₄MPT dehydrogenase (mtd), and formyl-methnaofuran:H₄MPT formyltransferase (ftr) were constructed by homologous recombination-mediated gene replacement as described previously (21). To do this, a plasmid that contained ca. 2 kb of the regions 5' and 3' of the gene of interest was constructed for each deletion. Between these regions was a pac-hpt cassette flanked by two Flp recombinase target (Frt) sites. Each plasmid was linearized and introduced into the parent strain, where a double recombination event between the upstream and downstream regions on the plasmid and the chromosome resulted in replacement of the chromosomal gene with the pac-hpt cassette. To remove the pac-hpt cassette and allow us to reuse puromycin as an antibiotic, the gene encoding Flp recombinase was introduced into each deletion strain on a nonreplicating plasmid, pMR55. The Flp recombinase is able to excise the DNA between the two Frt sites, thus removing the pac-hpt cassette and resulting in a strain that is puromycin sensitive and 8-ADP resistant and in which the gene of interest has been deleted. Deletion of the gene of interest and removal of the pac-hpt cassette in each strain were then verified by Southern blotting (data not shown).

Growth phenotypes of M. barkeri methyl oxidation deletion strains. The abilities of the mtr, mer, mtd, and ftr mutants to grow on a variety of substrates were determined (Table 4). While all of the mutants were able to grow using a combination of H₂/CO₂ and methanol, none of them were able to utilize methanol, acetate, or H₂/CO₂ alone as a growth substrate. These results suggest that none of the genes examined are required for growth via the methyl reduction pathway, whereas all of them are required for growth via the hydrogenotrophic, methylotrophic, and acetoclastic pathways. We previously showed that the mtr mutant is able to use a combination of methanol and acetate for growth (26). Because deletion of mer, mtd, or ftr should prevent oxidation of acetate to CO₂, we expected that the deletion mutants would not grow on methanol plus acetate, an expectation that was fulfilled by the mtd and ftr mutants. Surprisingly, the mer mutant was able to grow, albeit very slowly, on this combination. The doubling time for the mer mutant on methanol plus acetate was determined to be 130 ± 15 h, which is 10-fold longer than the doubling time observed for the mtr mutant (13 ± 1 h) (26).

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TABLE 4. Growth of M. barkeri deletion strains on various mediaa

Methane production in mutant cell suspensions. Resting cell suspensions of each strain were assayed for the production of CH₄ and CO₂ from various substrates (Table 5). Each of the mutants produced CH₄ from methanol plus H₂, and, as expected from the growth phenotype data, none of the mutants produced any CH₄ from H₂/CO₂ alone. Interestingly, both the mtr and mer mutants produced significant amounts of CH₄ and CO₂ from methanol alone, while the mtd and ftr mutants did not generate any CH₄ or CO₂ from methanol alone. Given that previous studies have demonstrated that a mch mutant is also unable to produce methane from methanol alone (11), these data strongly suggest that the methanol-oxidizing bypass pathway enters the standard C1 oxidation pathway at the level methylene-H₄MPT.

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TABLE 5. Methane and carbon dioxide produced from various substrates by M. barkeri cell suspensions

To demonstrate more directly that methanol was oxidized to CO₂ by the mtr and mer mutants but not by the mtd and ftr mutants, the cell suspension assays were repeated with [¹³C]methanol as the sole substrate. As shown in Table 6, both the mtr and mer mutants produced a significant amount of CH₄ and CO₂ from methanol, while the mtd and ftr mutants did not. Furthermore, all of the CH₄ and all of the CO₂ produced by the mtr and mer mutants were labeled, conclusively showing that these two mutants were able to bypass the steps catalyzed by Mtr and Mer and oxidize methanol to CO₂.

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TABLE 6. 13CH4 and 13CO2 production by M. barkeri deletion strain cell suspensions containing [13C]methanola

Biochemical assays for methanol dehydrogenase in C1 pathway mutants. Formaldehyde can condense spontaneously with H₄MPT to form methylene-H₄MPT (6, 16), which led us to speculate that formaldehyde might be an intermediate in the bypass pathway. If this is true, then methanol should first be oxidized to formaldehyde by a methanol dehydrogenase, as shown in Fig. 2. Accordingly, the oxidation of methanol to formaldehyde by mtr extracts was tested by monitoring the reduction of several different biological and artificial electron acceptors as described in Materials and Methods. Because the reaction is thermodynamically unfavorable in the oxidative direction with most of the electron acceptors tested, we also examined the reduction of formaldehyde to methanol with a variety of biological and artificial electrons donors. After exhaustive attempts, we were unable to detect any methanol dehydrogenase activity in mtr extracts, leaving the role of this enzymatic activity in question.

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FIG. 2. Proposed methyl oxidation bypass pathway. In the absence of Mtr or Mer (gray arrows), it is proposed that methanol is oxidized to formaldehyde via a methanol dehydrogenase (dashed arrows). Formaldehyde may then condense with H4MPT to form methylene-H4MPT. This reaction may occur spontaneously, or it may be catalyzed by the formaldehyde-activating enzyme (Fae). Methylene-H4MPT then enters the main oxidative branch of the methylotrophic pathway, where it can be further oxidized to CO2. For an explanation of abbreviations, see the legend to Fig. 1.

Involvement of the formaldehyde-activating enzyme (Fae) in the bypass pathway. In methylotrophic bacteria condensation of formaldehyde with H₄MPT to form methylene-H₄MPT is thought to play an important role in formaldehyde catabolism and detoxification. Although the spontaneous reaction occurs at a significant rate, the rate is not sufficient to keep up with the rate of C₁ metabolism (16). In Methylobacterium extorquens AM1 this problem is solved by the presence of the formaldehyde-activating enzyme (Fae), which catalyzes the formation of methylene-H₄MPT from free formaldehyde and H₄MPT (16, 24). The genome of M. barkeri Fusaro contains two fae homologues, faeA and faeB-hpsB, both of which have been purified and shown to catalyze the condensation of formaldehyde and H₄MPT to form methylene-H₄MPT in vitro (9). We therefore wanted to test whether the fae homologues found in the M. barkeri genome were required for the bypass pathway shown in Fig. 2.

We attempted to isolate faeA and faeB-hpsB single-deletion mutants of M. barkeri, as well as double-deletion mutants with the mtr and mer backgrounds. We were unable to obtain an faeB-hpsB deletion mutant, suggesting that this gene may be essential for growth in M. barkeri. We were able to construct mutants with a faeA deletion in M. barkeri and in the mtr and mer strains. Each faeA mutant was isolated while it was growing on methanol plus H₂/CO₂ and was confirmed by Southern blotting (data not shown). The faeA mutants were able to grow on all methanogenic substrates (Table 4). The double mutants were able to grow only on methanol plus H₂/CO₂, which is the same phenotype observed for the mtr and mer single-deletion strains. Interestingly, the faeA mtr mutant was able to grow on a combination of methanol plus acetate, while the faeA mer mutant was not.

To determine whether Fae plays a role in the bypass pathway, resting cell suspensions of the fae deletion strains were also assayed to determine the CH₄ and CO₂ production from methanol. As shown in Table 7, the mtr faeA and mer faeA double mutants produced total amounts of CH₄ and CO₂ similar to the amounts produced by the mtr and mer single mutants, and their rates of CH₄ production from methanol were similar to those of the mtr and mer single mutants. To focus more directly on the formaldehyde condensation reaction, we also examined the ability of fae cell extracts to form methyl-CoM from formaldehyde, CoM, and H₂. In this assay, Fae should catalyze the condensation of formaldehyde with H₄MPT that is present in the extract to form methylene-H₄MPT. Methylene-H₄MPT is then reduced to the methyl level with H₂ as a reductant and transferred to CoM, forming methyl-CoM (26). The rate of CoM consumption by fae extracts (23.6 nmol/min/mg of extract) was comparable to the rate of the wild-type strain (24.5 nmol/min/mg of extract). These data suggest that the fae strain was able to condense formaldehyde and H₄MPT to methylene-H₄MPT. (It should be noted, however, that an effect of the fae mutation would be missed if some other step in this multienzyme reaction is rate limiting.) Taken together, these data suggest that Fae does not play an obligate role in the bypass pathway, possibly because another activity is able to substitute for this enzyme. Our inability to isolate the faeB-hpsB deletion mutant suggests a logical candidate for the source of this activity.

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TABLE 7. Methane and carbon dioxide production by M. barkeri fae deletion strains supplemented with methanola

Role of bypass in utilization of methanol plus acetate in the mer mutant. We have previously demonstrated that the mtr mutant is able to use a combination of methanol plus acetate for growth (26). In this strain, both the methyl and carbonyl groups of acetate are oxidized to CO₂, producing reducing equivalents necessary to reduce methanol to CH₄. Unexpectedly, the mer mutant is also able to grow on a combination of methanol plus acetate, although at a much lower rate than the mtr strain. To determine how the mer mutant metabolizes methanol and acetate, one of the following substrate combinations was added to cell suspensions: [¹³C]methanol plus unlabeled acetate, methanol plus [1-¹³C]acetate, or methanol plus [2-¹³C]acetate. After incubation, production of ¹³CH₄ and ¹³CO₂ was measured using GC-MS. As Table 8 shows, mer cells were able to convert both methanol and acetate to CH₄ and CO₂. The majority of the CH₄ produced originated from methanol; 77% of the CH₄ generated came from methanol, and 26% came from the methyl group of acetate. None of the methyl groups of acetate were converted to CO₂, presumably because deletion of mer prevented the oxidation of the methyl groups derived from acetate via the methyl oxidation branch of the methylotrophic pathway. More importantly, 47% of the CO₂ produced came from methanol, indicating that a significant amount of flux occurred through the bypass pathway. In contrast, the mtr mutant oxidizes very little methanol (26). Thus, the mer mutant utilizes the bypass pathway at a much higher rate than the mtr mutant when the organisms are grown on a combination of methanol plus acetate. Coupled with the large difference in growth rates, these data indicate that growth on methanol plus acetate occurs via very different metabolic pathways in the mtr and mer mutants.

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TABLE 8. 13CH4 and 13CO2 production by M. barkeri mer and mtr cell suspensions grown on methanol plus acetate

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The data presented in this paper provide new insight into the roles of the C1 oxidation/reduction pathway in Methanosarcina. Although we have previously shown that mch mutations block both the methylotrophic and hydrogenotrophic pathways, it is now clear that this is a general feature of C1 pathway genes, including ftr, mch, mtd, mer, and the mtr operon. (Mutants lacking the formyl-methanofuran dehydrogenase [Fmd] have not been examined yet because four putative operons need to be deleted to construct an Fmd-deficient mutant [15]). Further, it is also clear that each of these C1 pathway genes is required for growth on acetate, while none is required for growth via the methyl reduction pathway. Nevertheless, there are clear phenotypic differences between many of the C1 pathway mutants. In particular, both the mer and mtr mutants are capable of oxidizing methanol to CO₂ via a bypass pathway, which is referred to here as the Mtr/Mer bypass. Further, both of these strains are able to grow using a combination of methanol and acetate, although our data suggest that this occurs via substantially different pathways.

In the acetoclastic pathway, acetate undergoes dismutation to CO₂ and CH₄, where the oxidation of the carbonyl group to CO₂ provides reducing equivalents necessary to reduce the methyl group to CH₄ (8). The carbonyl group is first cleaved and oxidized to CO₂ by the carbon monoxide dehydrogenase, while the methyl group is then transferred to H₄MPT (8). The methyl group thus enters the main methanogenic pathway at methyl-H₄MPT, where it is transferred to CoM by Mtr and then reduced to CH₄ (8). Thus, it appears that the C1 oxidation branch of the methylotrophic pathway is not required for methanogenesis from acetate and, accordingly, that mutations in this pathway should not prevent growth on acetate. However, it has been reported that a significant fraction of acetate methyl groups are oxidized to CO₂ during growth on acetate, most likely through the methyl oxidation branch of the methylotrophic pathway (14). This suggests that flux through the methyl oxidation branch is needed during acetoclastic growth in order to generate the reducing equivalents necessary for anabolic reactions during growth (12), which probably explains why none of the methyl oxidation deletion mutants characterized here are able to grow on acetate alone.

We also showed that both the mer and mtr mutants are capable of mixotrophic growth on a combination of acetate and methanol but use distinct pathways. We previously demonstrated that the mtr strain was able to utilize methanol plus acetate by using a unique combination of the methylotrophic and acetoclastic pathways (26). In this strain, oxidation of both the carbonyl and methyl groups of acetate is needed to drive the reduction of methanol. Because Mer is required for the oxidation of the acetate methyl group to CO₂, this pathway cannot be available to the mer mutant. However, as described above, aceticlastic methanogenesis is not blocked in the C1 pathway mutants; rather, these mutants cannot generate the reducing equivalents needed for biosynthesis. Our data show that when the mer mutant is given methanol in combination with acetate, a significant fraction of the methanol is oxidized via the bypass pathway. This flux seems to be sufficient to overcome any biosynthetic block that prevents growth on acetate alone. We therefore propose that the mer mutant utilizes methanol and acetate via a novel pathway that requires the Mtr/Mer bypass pathway, as shown in Fig. 3.

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FIG. 3. Proposed cometabolism of methanol and acetate by the mer mutant. mer cells are able to utilize the proposed bypass pathway to grow on a combination of methanol plus acetate. In this pathway, methanol is oxidized to CO2 via the bypass pathway (dashed arrows) and is reduced to CH4 via the methyl reduction portion of the methylotrophic pathway. Acetate is then dismutated to CO2 and CH4, with the carbonyl group of acetate oxidized to CO2 and the methyl group of acetate reduced to CH4. Flux of the methyl group of acetate through the methyl oxidation branch is prevented by deletion of mer, and therefore, mer cells are unable to grow on acetate alone. Ac, acetate. For an explanation of other abbreviations, see the legend to Fig. 1.

Our data clearly show that the Mtr/Mer bypass pathway enters the main C1 oxidation pathway at the methylene level. However, we were unable to provide biochemical evidence for the production of free formaldehyde. This result is consistent with previous studies that failed to detect methanol dehydrogenase in M. barkeri (13) and the absence of homologs of genes encoding known methanol dehydrogenases, including glutathione-dependent dehydrogenases, in the genome sequence (3). Nonetheless, there are several short-chain alcohol dehydrogenases annotated in the M. barkeri Fusaro genome that could possibly oxidize methanol to formaldehyde (Mbar_A0771, Mbar_A0783, Mbar_A0784, and Mbar_A2344) (15).

There are several possible reasons why no methanol dehydrogenase activity was detected in mtr extracts. The assays that we performed may have failed due to an inability to identify an appropriate electron acceptor. Given the low redox potentials of the methanogenic electron carriers (for F₄₂₀, E^o' = –360 mV; for ferredoxin, E^o' = –410 mV; and for methanophenazine, E^o' = –255 mV) relative to that of the methanol-formaldehyde couple (E^o' = –185 mV), it seems unlikely that these electron carriers would function in accepting electrons from methanol. Higher-potential acceptors, such as quinones, have not been reported for Methanosarcina, nor did they function in our assays. Alternatively, the inability to detect methanol dehydrogenase activity could have been due to a lack of expression of the methanol dehydrogenase in the mtr mutant under the growth conditions that we used. For these assays, mtr cells were grown on methanol plus acetate. Although there is some flux of methyl groups through the bypass pathway (Table 8), it does not seem to be required for growth of the mtr mutant on this combination of substrates. Therefore, a strain that requires the bypass pathway for growth might express higher levels of methanol dehydrogenase, making detection in extracts much easier. Because the mer strain requires the bypass pathway for growth on methanol plus acetate, this strain is perhaps a better candidate for detection of methanol dehydrogenase activity.

Genetic approaches also failed to provide evidence for the involvement of free formaldehyde. The abilities of the faeA mutant that we constructed to oxidize methanol to CO₂ and to catalyze the in vitro conversion of formaldehyde, CoM, and H₂ gas to methyl-CoM were unimpaired. It seems very likely, however, that this was due to the presence of a second copy of Fae in M. barkeri, encoded by the faeB-hpsB gene. The faeB-hpsB gene from M. barkeri has been overexpressed in E. coli, and cell extracts of the recombinant E. coli strain have been shown to catalyze the condensation of formaldehyde and H₄MPT to methylene-H₄MPT (9). Our inability to delete faeB-hpsB in M. barkeri led us to assume that FaeB-HpsB is essential for growth in M. barkeri and supports the importance of this condensation reaction in the metabolism of M. barkeri.

Finally, it remains unclear why the Mtr/Mer bypass pathway does not allow growth on methanol alone. Because our data suggest that formaldehyde may be an intermediate in this bypass pathway, one could imagine that formaldehyde toxicity may play a role in the lack of growth via the bypass pathway. In order to support growth, methanol oxidation to formaldehyde must occur at a high rate. If the formaldehyde generated is not efficiently converted to methylene-H₄MPT, then it is possible that the accumulation of this highly reactive compound could be toxic for the cells. However, this does not occur in the mer mutant during growth on methanol plus acetate, which does not support this argument. It is also possible that the proteins involved in the bypass pathway, specifically a methanol dehydrogenase and a formaldehyde-condensing enzyme, are inefficient and unable to convert the methyl groups to CO₂ at a rate high enough to allow growth via methanol-dependent methanogenesis. One would presume, however, that bypassing the energy-consuming Mtr methyl transfer would allow the organism to generate more ATP and would thus result in better growth through the bypass on methanol. We previously argued that perhaps there is simply not sufficient energy available from methanol disproportionation through the bypass pathway to support extra ATP generation (26). Although the bypass pathway proposed in this study does not allow growth on methanol alone, it is sufficient to allow growth of the mer mutant on methanol in combination with acetate. Thus, this bypass pathway is able to play a physiological role in the cell. Further studies are needed to identify and characterize the proteins involved in this pathway to determine what physiological role they may play in the cell and hopefully reveal why the bypass pathway is unable to support growth on methanol alone.

 
We thank L. Daniels for providing F₄₂₀ and N. Buan, G. Kulkarni, and A. Bose for critical reviews of the manuscript.

This work was supported by National Science Foundation grant MCB0517419 to W.W.M. and by a National Science Foundation predoctoral fellowship to P.V.W.

The opinions, findings, conclusions, and recommendations expressed in this paper are those of the authors and do not necessarily reflect the views of the National Science Foundation.

 
* Corresponding author. Mailing address: Department of Microbiology, University of Illinois at Urbana-Champaign, 601 S. Goodwin Avenue, Urbana, IL 61801. Phone: (217) 244-1943. Fax: (217) 244-6697. E-mail: metcalf{at}uiuc.edu

Published ahead of print on 4 January 2008.

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

 

1
1. Ausubel, F. M., R. Brent, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A. Smith, and K. Struhl. 1992. Current protocols in molecular biology, vol. 1 and 2. John Wiley & Sons, New York, NY.
2
2. Boccazzi, P., J. K. Zhang, and W. W. Metcalf. 2000. Generation of dominant selectable markers for resistance to pseudomonic acid by cloning and mutagenesis of the ileS gene from the archaeon Methanosarcina barkeri Fusaro. J. Bacteriol. 182:2611-2618.[Abstract/Free Full Text]
3
3. Daussmann, T., A. Aivasidis, and C. Wandrey. 1997. Purification and characterization of an alcohol:N,N-dimethyl-4-nitrosoaniline oxidoreductase from the methanogen Methanosarcina barkeri DSM 804 strain Fusaro. Eur. J. Biochem. 248:889-896.[Medline]
4
4. Deppenmeier, U. 2002. The unique biochemistry of methanogenesis. Prog. Nucleic Acid Res. Mol. Biol. 71:223-283.[Medline]
5
5. Deppenmeier, U., V. Muller, and G. Gottschalk. 1996. Pathways of energy conservation in methanogenic archaea. Arch. Microbiol. 165:149-163.[CrossRef]
6
6. Escalante-Semerena, J. C., and R. S. Wolfe. 1984. Formaldehyde oxidation and methanogenesis. J. Bacteriol. 158:721-726.[Abstract/Free Full Text]
7
7. Ferry, J. G. 1999. Enzymology of one-carbon metabolism in methanogenic pathways. FEMS Microbiol. Rev. 23:13-38.[CrossRef][Medline]
8
8. Ferry, J. G. 1993. In J. G. Ferry (ed.), Methanogenesis: ecology, physiology, biochemistry and genetics, p. 304-334. Chapman & Hall, New York, NY.
9
9. Goenrich, M., R. K. Thauer, H. Yurimoto, and N. Kato. 2005. Formaldehyde activating enzyme (Fae) and hexulose-6-phosphate synthase (Hps) in Methanosarcina barkeri: a possible function in ribose-5-phosphate biosynthesis. Arch. Microbiol. 184:41-48.[CrossRef][Medline]
10
10. Gottschalk, G., and R. K. Thauer. 2001. The Na⁺-translocating methyltransferase complex from methanogenic archaea. Biochim. Biophys. Acta 1505:28-36.[Medline]
11
11. Guss, A. M., B. Mukhopadhyay, J. K. Zhang, and W. W. Metcalf. 2005. Genetic analysis of mch mutants in two Methanosarcina species demonstrates multiple roles for the methanopterin-dependent C-1 oxidation/reduction pathway and differences in H₂ metabolism between closely related species. Mol. Microbiol. 55:1671-1680.[CrossRef][Medline]
12
12. Jablonski, P. E., A. A. DiMarco, T. A. Bobik, M. C. Cabell, and J. G. Ferry. 1990. Protein content and enzyme activities in methanol- and acetate-grown Methanosarcina thermophila. J. Bacteriol. 172:1271-1275.[Abstract/Free Full Text]
13
13. Keltjens, J. T., and G. D. Vogels. 1993. Conversion of methanol and methylamines to methane and carbon dioxide, p. 253-303. In J. G. Ferry (ed.), Methanogenesis: ecology, physiology, biochemistry and genetics. Chapman & Hall, New York, NY.
14
14. Krzycki, J. A., R. H. Wolkin, and J. G. Zeikus. 1982. Comparison of unitrophic and mixotrophic substrate metabolism by acetate-adapted strain of Methanosarcina barkeri. J. Bacteriol. 149:247-254.[Abstract/Free Full Text]
15
15. Maeder, D. L., I. Anderson, T. S. Brettin, D. C. Bruce, P. Gilna, C. S. Han, A. Lapidus, W. W. Metcalf, E. Saunders, R. Tapia, and K. R. Sowers. 2006. The Methanosarcina barkeri genome: comparative analysis with Methanosarcina acetivorans and Methanosarcina mazei reveals extensive rearrangement within methanosarcinal genomes. J. Bacteriol. 188:7922-7931.[Abstract/Free Full Text]
16
16. Marx, C. J., L. Chistoserdova, and M. E. Lidstrom. 2003. Formaldehyde-detoxifying role of the tetrahydromethanopterin-linked pathway in Methylobacterium extorquens AM1. J. Bacteriol. 185:7160-7168.[Abstract/Free Full Text]
17
17. Metcalf, W. W., J. K. Zhang, E. Apolinario, K. R. Sowers, and R. S. Wolfe. 1997. A genetic system for Archaea of the genus Methanosarcina: liposome-mediated transformation and construction of shuttle vectors. Proc. Natl. Acad. Sci. USA 94:2626-2631.[Abstract/Free Full Text]
18
18. Metcalf, W. W., J. K. Zhang, X. Shi, and R. S. Wolfe. 1996. Molecular, genetic, and biochemical characterization of the serC gene of Methanosarcina barkeri Fusaro. J. Bacteriol. 178:5797-5802.[Abstract/Free Full Text]
19
19. Metcalf, W. W., J. K. Zhang, and R. S. Wolfe. 1998. An anaerobic, intrachamber incubator for growth of Methanosarcina spp. on methanol-containing solid media. Appl. Environ. Microbiol. 64:768-770.[Abstract/Free Full Text]
20
20. Pritchett, M. A., J. K. Zhang, and W. W. Metcalf. 2004. Development of a markerless genetic exchange method for Methanosarcina acetivorans C2A and its use in construction of new genetic tools for methanogenic archaea. Appl. Environ. Microbiol. 70:1425-1433.[Abstract/Free Full Text]
21
21. Rother, M., and W. W. Metcalf. 2005. Genetic technologies for Archaea. Curr. Opin. Microbiol. 8:745-751.[Medline]
22
22. Sowers, K. R., J. E. Boone, and R. P. Gunsalus. 1993. Disaggregation of Methanosarcina spp. and growth as single cells at elevated osmolarity. Appl. Environ. Microbiol. 59:3832-3839.[Abstract/Free Full Text]
23
23. Thauer, R. K. 1998. Biochemistry of methanogenesis: a tribute to Marjory Stephenson. Microbiology 144:2377-2406.[Abstract/Free Full Text]
24
24. Vorholt, J. A., C. J. Marx, M. E. Lidstrom, and R. K. Thauer. 2000. Novel formaldehyde-activating enzyme in Methylobacterium extorquens AM1 required for growth on methanol. J. Bacteriol. 182:6645-6650.[Abstract/Free Full Text]
25
25. Wanner, B. L. 1986. Novel regulatory mutants of the phosphate regulon in Escherichia coli K-12. J. Mol. Biol. 191:39-58.[CrossRef][Medline]
26
26. Welander, P. V., and W. W. Metcalf. 2005. Loss of the mtr operon in Methanosarcina blocks growth on methanol, but not methanogenesis, and reveals an unknown methanogenic pathway. Proc. Natl. Acad. Sci. USA 102:10664-10669.[Abstract/Free Full Text]

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Journal of Bacteriology, March 2008, p. 1928-1936, Vol. 190, No. 6
0021-9193/08/$08.00+0     doi:10.1128/JB.01424-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

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