• We are sorry, but NCBI web applications do not support your browser and may not function properly. More information
Logo of pnasPNASInfo for AuthorsSubscriptionsAboutThis Article
Proc Natl Acad Sci U S A. Nov 21, 2006; 103(47): 17921–17926.
Published online Nov 13, 2006. doi:  10.1073/pnas.0608833103
PMCID: PMC1693848
Microbiology

An unconventional pathway for reduction of CO2 to methane in CO-grown Methanosarcina acetivorans revealed by proteomics

Abstract

Methanosarcina acetivorans produces acetate, formate, and methane when cultured with CO as the growth substrate [Rother M, Metcalf WW (2004) Proc Natl Acad Sci USA 101:16929–16934], which suggests novel features of CO metabolism. Here we present a genome-wide proteomic approach to identify and quantify proteins differentially abundant in response to growth on CO versus methanol or acetate. The results indicate that oxidation of CO to CO2 supplies electrons for reduction of CO2 to a methyl group by steps and enzymes of the pathway for CO2 reduction determined for other methane-producing species. However, proteomic and quantitative RT-PCR results suggest that reduction of the methyl group to methane involves novel methyltransferases and a coenzyme F420H2:heterodisulfide oxidoreductase system that generates a proton gradient for ATP synthesis not previously described for pathways reducing CO2 to methane. Biochemical assays support a role for the oxidoreductase, and transcriptional mapping identified an unusual operon structure encoding the oxidoreductase. The proteomic results further indicate that acetate is synthesized from the methyl group and CO by a reversal of initial steps in the pathway for conversion of acetate to methane that yields ATP by substrate level phosphorylation. The results indicate that M. acetivorans utilizes a pathway distinct from all known CO2 reduction pathways for methane formation that reflects an adaptation to the marine environment. Finally, the pathway supports the basis for a recently proposed primitive CO-dependent energy-conservation cycle that drove and directed the early evolution of life on Earth.

Keywords: anaerobic, Archaea, carbon monoxide

Carbon monoxide (CO), an atmospheric pollutant that binds tightly to hemoglobin, is held below toxic levels in part by both aerobic and anaerobic microbes (1). The microbial metabolism of CO is an important component of the global carbon cycle (1, 2), and CO is believed to have been present in the atmosphere of early Earth that fueled the evolution of primitive metabolisms (37). Investigations of aerobic species from the Bacteria domain have contributed important insights into microbial CO oxidation (8, 9), as have investigations of anaerobes from the Bacteria domain that conserve energy by coupling CO oxidation to H2 evolution (1012). Further understanding has been derived from studies of CO-using anaerobes from the Bacteria domain that conserve energy by oxidizing CO and reducing CO2 to acetate (13, 14) or reducing sulfate to sulfide (15). Far less is known for pathways of the few CO-using species in the Archaea domain that have been described. Methanothermobacter thermautotrophicus, Methanosarcina barkeri, and Methanosarcina acetivorans obtain energy for growth by converting CO to methane (1620). Although methane formation from CO first was reported in 1947 (21), a comprehensive understanding of the overall pathway for any species has not been reported. It is postulated that M. barkeri oxidizes CO to H2, and the H2 is reoxidized to provide electrons for reducing CO2 to methane (16). It is postulated further that H2 production is essential for ATP synthesis during growth on CO (16, 22, 23). M. acetivorans was isolated from marine sediments where giant kelp is decomposed to methane (24). The flotation bladders of kelp contain CO that is a presumed substrate for M. acetivorans in nature. M. acetivorans produces acetate and formate in addition to methane during CO-dependent growth (17), the first report of any product in addition to methane and CO2 during growth of methane-producing species. Further, H2 was not detected during growth on CO (17). These unorthodox characteristics suggest that the pathway for metabolism of CO and mechanisms for energy conservation by M. acetivorans are unique among methane-producing Archaea. Here we describe a global proteomic analysis of CO metabolism in M. acetivorans. The results suggest that the pathway for methane formation involves novel methyltransferases and a mechanism for energy conservation not previously reported for CO2-reducing pathways. Finally, the results provide support for the basis of a recent proposal that acetate formation from CO in M. acetivorans is the vestige of a primitive energy-conservation cycle that drove and directed the early evolution of life on Earth (7).

Results and Discussion

Fig. 1 shows the time course obtained for CO-dependent growth of M. acetivorans. Acetate, formate, methane, and CO2 were end products, and H2 was not detected (limit of detection = 0.008%). These results are consistent with the previous report for CO-dependent growth of M. acetivorans (17) except that production of methane significantly exceeded acetate throughout growth and only minor amounts of formate was produced (Fig. 1). The after-growth carbon balance indicated that no other reduced products were synthesized. These results indicate that CO2 reduction to methane plays a prominent role in supporting CO-dependent growth of M. acetivorans.

Fig. 1.
Time course of growth by Methanosarcina acetivorans C2A with CO as the sole source of carbon and energy. Each point is the mean of five replicate cultures. ●, CO2; [filled triangle], CO; ■, CH4; □, acetate; [open triangle], A660; ◊, ...

A high-sensitivity liquid chromatography/mass spectrometry analysis, using a hybrid linear ion trap/Fourier-transform ion cyclotron resonance (LTQ/FTICR)-MS, was used to identify, quantify, and compare individual proteins from CO-grown cells metabolically labeled with 14N with separately methanol-grown and acetate-grown cells labeled with 15N. Abundance ratios of 1,023 proteins were obtained for CO versus methanol grown cells and 846 proteins for CO versus acetate grown cells, corresponding to a total of 1,125 unique proteins (Table 4, which is published as supporting information on the PNAS web site), representing 25% of the 4,524 genes reported in the genome (25). Of these unique proteins, 99 were found to have ≥3-fold differential abundance between CO- and methanol-grown cells and 176 between CO- and acetate-grown cells. Of the 1,125 proteins in Table 4, many were annotated for functions in pathways for methane formation from the dismutation of methanol, reduction of CO2, and fermentation of acetate (Table 1). The function of these proteins and their differential abundance (Table 1) lead to the pathway for conversion of CO to acetate and methane shown in Fig. 2. No evidence has been reported for the mechanism of formate production (17), and the proteomic analyses reported here yield no clues as to the mechanism; thus, a pathway for production of formate is not shown in Fig. 2.

Fig. 2.
Pathway proposed for the conversion of CO to acetate and methane by M. acetivorans. Proposed enzymes catalyzing steps 1–13 are indicated in Table 1. Fdo, oxidized ferredoxin; Fdr, reduced ferredoxin; F420, coenzyme F420; MF, methanofuran; THMPT, ...
Table 1.
Relative abundance of selected proteins in CO-grown versus methanol-grown and CO-grown versus acetate-grown M. acetivorans strain C2A

In the pathway shown in Fig. 2, CO2 is reduced to methyl-tetrahydromethanopterin (THMPT) in steps 2–6 catalyzed by paralogs of enzymes characterized from CO2-reducing methanoarchaea (26). Enzymes catalyzing steps 2, 3, 5, and 6 were at least 10-fold more abundant in CO-grown versus acetate-grown M. acetivorans, and the enzyme catalyzing step 4 was 2-fold greater (Table 1). These steps and enzymes are not involved in acetate fermentation to methane (27); thus, the results suggest these enzymes are differentially abundant in response to CO and support roles in steps 2–6. Further, M. acetivorans, like other Methanosarcina species, utilizes a reversal of steps 2–6 in the pathway of methanol dismutation to CO2 and CH4 (28, 29); thus, the finding that each of the five enzymes was at approximately the same levels in CO-grown versus methanol-grown cells further supports roles in steps 2–6 (Fig. 2). Subunits of both the molybdenum (MA0304–MA0309) and tungsten (MA0832–MA0835) forms of formyl-methanofuran (MF) dehydrogenase catalyzing step 2 were substantially elevated in CO-grown versus acetate-grown cells, a result suggesting both metal forms function during growth on CO. However, the relative levels between CO- and methanol- or acetate-grown cells suggest that the tungsten form is favored during growth on CO.

Pathways for methanogenesis from all substrates involve an eight-subunit methyl-THMPT:HS-coenzyme M (CoM) methyl transferase (MtrABCDEFGH) complex that contains a methyl-accepting corrinoid cofactor (30). However, several subunits of the Mtr complex encoded by MA0269–MA0276 were detected with a 9-fold mean lower abundance in CO-grown versus acetate-grown M. acetivorans (Table 1), suggesting down-regulation of the complex in response to growth with CO that is supported by quantitative RT-PCR results obtained for expression of MA0269, MA0272, and MA0276 (Table 2). These results were in contrast to the relative abundances of McrA (MA4546) and McrG (MA4547) from the three-subunit CH3-S-CoM methyl reductase (McrABG) that is essential to all methanogenic pathways. Both McrA and McrG were approximately equal in abundance in CO-grown versus acetate- or methanol-grown cells (Table 1), a result confirming the expected role in catalyzing step 8 (Fig. 2) of the pathway for methanogenesis. Thus, the proteomic and RT-PCR results obtained for the Mtr subunits suggest a substantially diminished role for the Mtr complex in catalyzing step 7 (Fig. 2) compared with all other pathways for methanogenesis. Although unable to grow on methanol, a mutant of M. barkeri disrupted in the mtr operon is able to produce methane from methanol (31), providing experimental evidence for an uncharacterized alternative to the Mtr complex. Thus, it was interesting that the products of MA0859 and MA4384 were robustly elevated in CO-grown versus methanol-grown and CO-grown versus acetate-grown M. acetivorans, respectively (Table 1). Based on sequence analysis, these genes are predicted to encode putative methyl transferases (25) and are annotated as corrinoid proteins (www.tigr.org) consistent with this function. Quantitative RT-PCR analyses (Table 2) showed MA0859 and MA4384 substantially up-regulated in CO-grown versus acetate-grown cells. MA4558, encoding a third putative methyl transferase (25) and annotated as encoding a corrinoid protein (www.tigr.org), was up-regulated in CO-grown versus acetate-grown cells (Table 2). Further, the products of MA0859, MA4384, and MA4558 have >55% sequence identity to each other, and all three putative methyl transferases contain a C-terminal domain not found in other corrinoid-containing proteins. These results encourage investigation to determine whether the products of MA0859, MA4384, and MA4558 function in step 7 (Fig. 2).

Table 2.
Expression ratios of genes in acetate-grown versus CO-grown M. acetivorans determined by quantitative RT-PCR

The genome is annotated (www.tigr.org) with several CO dehydrogenases that have the potential to catalyze step 1a, providing CO2 and electrons for steps 2, 5, 6, 8, 9, and 10 in the pathway for methanogenesis (Fig. 2). MA1309 and MA3282 are annotated as encoding CooS, the CO dehydrogenase first described in Rhodospirillum rubrum (32). The products of MA1309 and MA3282 were not detected in CO-grown cells (Table 4), although quantitative RT-PCR analysis (Table 2) indicated modest up-regulation of MA1309 in CO-grown versus acetate-grown cells consistent with a role for this CO dehydrogenase in the oxidation of CO. Although the CooS paralog in R. rubrum reduces ferredoxin (33), the enzyme in M. acetivorans has not been investigated, and the electron acceptor is unknown. The genome also is annotated (www.tigr.org) with duplicate gene clusters (MA1011–MA1016 and MA3860–MA3865), each encoding paralogs of a five-subunit CO dehydrogenase/acetyl-CoA synthase (CdhABCDE) complex first described in Methanosarcina thermophila (34, 35). Three subunits (MA1014–MA1016 and MA3860–MA3862) from each complex were detected and found to be elevated an average of 20-fold in CO-grown versus methanol-grown cells in contrast to an ≈2-fold differential abundance versus acetate-grown cells (Table 1). The Cdh complex is not involved in the pathway for conversion of methanol to methane but is central to the pathway for conversion of acetate to methane of Methanosarcina species (26); thus, the relative levels in CO-grown versus methanol- or acetate-grown M. acetivorans suggest a role for the duplicate CdhABCDE complexes during growth on CO. Although the CdhABCDE complex of Methanosarcina species oxidizes CO and reduces ferredoxin (36, 37), consistent with a role in steps 1a and 1b (Fig. 2), a more certain function is catalysis of step 11 as described below.

Regardless which CO dehydrogenase functions in step 1a (Fig. 2), reduction of ferredoxin (step 1b) is most likely essential when considering ferredoxin has been proposed as the direct electron donor for formyl-MF dehydrogenases (38, 39) (step 2). Further, reduced coenzyme F420 is the direct electron donor for characterized enzymes (26) catalyzing steps 5 and 6 (Fig. 2); thus, a mechanism for reducing F420 also is expected (step 1c). Characterization of the CooS CO dehydrogenase from any methanogen has not been reported, and the electron acceptor is unknown. Although ferredoxin is the electron acceptor for CooS from Rhodospirillum rubrum (33), reduction of F420 by the CooS homolog from M. acetivorans cannot be ruled out. None of the proteins detected in the proteome of CO-grown M. acetivorans (Tables 1 and 4) have the potential to oxidize ferredoxin and reduce F420, and the mechanism by which F420 is reduced is unknown.

In all known methanogenic pathways, the reduction of CH3-S-CoM with coenzyme B (HS-CoB) (step 8) produces the heterodisulfide of CoM-S-S-CoB, which requires reduction of the disulfide bond by a heterodisulfide reductase (step 9) to regenerate the sulfhydryl forms of the cofactors (Fig. 2). Both subunits of the HdrDE heterodisulfide reductase (MA0688 and MA0687) were present in CO-grown cells, which approximated the amounts in methanol- or acetate-grown cells (Table 1). HdrDE functions in the pathway for conversion of methanol to methane in all Methanosarcina species investigated (40) and is proposed to function in the pathway for conversion of acetate to methane in M. thermophila (41) and M. barkeri (42). Thus, the relative levels of HdrDE in CO-grown versus acetate- or methanol-grown cells (Table 1) support a role for HdrDE in step 9 of the pathway for conversion of CO to methane by M. acetivorans (Fig. 2). In obligate CO2-reducing methanogens, reduction of CoM-S-S-CoB is accomplished by the HdrABC-type of heterodisulfide reductase, and electrons are supplied to it from H2 catalyzed by the MvhAGD hydrogenase (43). M. acetivorans is unable to grow by reducing CO2 to methane with H2 (24), and genes encoding the MvhAGD hydrogenase are absent from the genome (www.tigr.org). Thus, a major role for HdrABC is improbable during growth with CO.

The electron donor to the membrane-bound HdrDE in all Methanosarcina species previously investigated is methanophenazine, a membrane-bound quinone-like electron carrier that pumps protons to the outside upon reduction and oxidation (40). In the methanol pathway, methanophenazine transfers electrons from the F420H2 dehydrogenase complex (FpoABCDHIJKLMNO) to HdrDE (40). The 12-subunit Fpo complex is membrane-bound and also functions as a proton pump. The genome of M. acetivorans is annotated with a gene cluster (MA1495–MA1507) encoding a Fpo complex for which the products of 11 subunits were found in CO-grown M. acetivorans at approximately the same levels as in methanol-grown cells (Table 1). Furthermore, the amounts were considerably more abundant in CO-grown versus acetate-grown cells where the Fpo complex does not function (27). F420H2:CoB-S-S-CoM oxidoreductase activity in cell extracts of CO-grown M. acetivorans was approximately 2- and 10-fold greater than for methanol- and acetate-grown cells, respectively (Table 3). Although unprecedented in pathways for the reduction of CO2 to methane, these results support a role for the Fpo complex, methanophenazine, and HdrDE in generation of a proton gradient that drives ATP synthesis in the pathway for CO-dependent reduction of CO2 to methane by M. acetivorans (Fig. 2, step 10). Transcriptional mapping indicated that the Fpo complex is encoded in an operon of 14 genes (Fig. 3A) containing duplicate fpoJ genes and an additional ORF (MA1494) annotated as a predicted protein (www.tigr.org), unique to M. acetivorans, that we designate here as fpoP (Fig. 3A). The sequence upstream of MA1494 contained a typical archaeal TATA-box (Fig. 3B), and a transcription start site was identified 22 nucleotides downstream of the TATA-box. Quantitative RT-PCR showed that MA1494 is up-regulated in CO-grown versus acetate-grown cells consistent with the differential abundances of other Fpo subunits encoded in the operon (Table 1). These results suggest that the Fpo complex of CO-grown M. acetivorans may have an unique subunit composition compared with other characterized Fpo complexes.

Fig. 3.
Transcriptional mapping of the fpoPABCDHIJJKLMNO operon of M. acetivorans. (A) Cotranscription determined by RT-PCR. Uppercase letters refer to subunits encoded by the genes represented by arrows showing the direction of transcription. Predicted RT-PCR ...
Table 3.
Coenzyme F420H2:CoB-S-S-CoM oxidoreductase activity in cell extracts of CO-, methanol-, and acetate-grown M. acetivorans

As discussed above, the relative abundances of subunits from two CdhABCDE complexes in CO-grown versus methanol- and acetate-grown M. acetivorans (Table 1) suggest a role during growth on CO. Although oxidation of CO to CO2 is one potential role, the most probable function is synthesis of acetyl-CoA from methyl-THMPT, CO, and CoA in step 11 (Fig. 2). The first CdhABCDE complex to be described was isolated from acetate-grown M. thermophila where it functions to cleave acetyl-CoA (34), the reverse of step 11 (Fig. 2). However, the reported synthesis of acetyl-CoA from CO, CH3I, and CoA by the Cdh complex from M. thermophila (44) supports a role for the Cdh complex as shown in step 11 of the pathway for CO conversion to acetate by M. acetivorans (Fig. 2). The duplicate CdhABCDE complexes encoded by MA1011–MA1016 and MA3860–MA3865 of M. acetivorans share >90% amino acid sequence identity (www.tigr.org), consistent with a similar function. A remarkable feature of the genomic sequence of M. acetivorans and Methanosarcina mazei is the extensive gene redundancy that has raised questions regarding the expression of duplicated genes (25, 45). As discussed above, the differential abundances for subunits of the duplicate CdhABCDE complexes in CO-grown versus methanol- or acetate-grown M. acetivorans (Table 1) indicate that both complexes are present in high amounts in CO-grown cells.

A role for phosphotransacetylase (MA3607) and acetate kinase (MA3606) in the pathway for conversion of CO to acetate (Fig. 2, steps 12 and 13) was demonstrated previously (17). Both enzymes were ≈4-fold less abundant in CO-grown versus acetate-grown cells (Table 1); however, 50- and 18-fold greater amounts of acetate kinase and phosphotransacetylase are reported for acetate-grown versus methanol-grown M. acetivorans (27), suggesting substantial amounts of both enzymes in acetate-grown cells. Thus, although lower amounts were detected in CO-grown versus acetate grown cells (Table 1), the levels of acetate kinase and phosphotransacetylase have the potential to supply adequate amounts for synthesis of acetate from acetyl-CoA during growth on CO. These results are consistent with a previous report (17) and suggest that the production of acetate yields ATP via a substrate level phosphorylation. Thus, ATP is synthesized by substrate level phosphorylation (Fig. 2, step 13) and driven by a proton gradient coupled to electron transport from F420H2 to CoB-S-S-CoM (Fig. 2, steps 9 and 10). The pathway in Fig. 2 provides support for the basis of the recent proposal that CO conversion to acetate in methanogens is the vestige of a primitive energy conservation cycle that was the dominant force that drove and directed the early evolution of life, including methanogenic pathways (7).

It has been proposed that CO-dependent growth of methanogens other than M. acetivorans involves oxidation of CO to H2 and reoxidation of the H2 to supply electrons for the reduction of CO2 to methane (16). The pathway shown in Fig. 2 is independent of H2 and consistent with the report that H2 is not a metabolite during CO-dependent growth of M. acetivorans (17). The only mechanism reported for energy conservation in the CO2-reduction pathway for methanogenesis is the reduction of CoB-S-S-CoM with H2, which generates a proton gradient driving ATP synthesis (46). The H2:CoB-S-S-CoM oxidoreductase system is composed of the MvhAGD hydrogenase (43) and the HdrABC-type of heterodisulfide reductase (47). As previously discussed, the evidence presented here suggests neither enzyme functions in the pathway for CO2 reduction to methane during CO-dependent growth of M. acetivorans. Instead, it appears M. acetivorans has adopted a H2-independent mechanism for energy conservation from the pathway for conversion of methanol to methane wherein the F420H2:CoB-S-S-CoM system shown in Fig. 2 (steps 9 and 10) also functions to pump protons for energy conservation. However, based on the transcriptional analysis presented above, it is tempting to speculate that adoption of the F420H2:CoB-S-S-CoM system for growth on CO may have required the addition of another subunit encoded by MA1494. The inability of M. acetivorans to metabolize H2 suggests that CO-dependent reduction of CO2 to formyl-MF (Fig. 2, step 2) also is different from previously characterized CO2-reduction pathways of methanogens. The oxidation of H2 and reduction of CO2 to produce formyl-MF is endergonic, and it has been shown that M. barkeri employs the Ech hydrogenase complex to catalyze the oxidation of H2 that is driven by an ion gradient (39, 48). The genome of M. acetivorans does not encode a functional Ech hydrogenase (25) (www.tigr.org); however, the reduction of CO2 to formyl-MF with CO as the electron donor is exergonic and does not require H2 and the Ech hydrogenase to drive the reaction (38). The H2-independent pathway in Fig. 2 also is consistent with the marine environment from which M. acetivorans was isolated (24). In marine environments sulfate-reducing microbes outcompete methanogens for H2 (49); thus, if H2 was an intermediate in metabolism, it could be lost to sulfate reducers. Indeed, it was shown recently that the electron transport chain in acetate-grown M. acetivorans does not involve H2 (27) in contrast to acetate-grown freshwater Methanosarcina species for which it is proposed that H2 and the Ech hydrogenase plays an essential role in electron transport and energy conservation (39, 48).

Conclusions

A highly sensitive proteomic analysis has identified proteins differentially abundant in CO-grown versus methanol- and acetate-grown M. acetivorans, revealing steps and enzymes in the decidedly unusual pathway of CO conversion to acetate and methane. Although the pathway involves reduction of CO2 to CH3-THMPT, similar to known CO2-reduction pathways, subsequent reduction of the methyl group from CH3-THMPT to methane appears to involve novel methyltransferases and a mechanism for energy conservation different from known pathways for CO2 reduction to methane. This unusual pathway is consistent with the idea that marine methanogens have evolved pathways independent of H2. Finally, the results support the basis for a recent proposal that the pathway for acetate formation from CO is the vestige of an ancient energy-conservation mechanism that directed the early evolution of life including the CO2-reduction and acetate fermentation pathways for methanogenesis.

Materials and Methods

Growth of M. acetivorans and Preparation of Samples for MS Analysis.

M. acetivorans C2A (DSM 804) was grown in high-salt media (50) with one of three growth substrates: (i) 1.0 atm of CO (1 atm = 101.3 kPa), (ii) 100 mM acetate, or (iii) 250 mM methanol. For the methanol and acetate-grown cultures, 14NH4Cl was substituted with 15NH4Cl (98%) (Sigma, St. Louis, MO). Cells were harvested in the midexponential phase of growth at an A660 of 0.3 (CO-grown), 0.8 (acetate-grown), and 0.6 (methanol-grown) as previously described (28, 29). Headspace gases and acetate were quantified by gas chromatographical analysis as described in ref. 51. Formate quantification was performed as described in ref. 52.

Protein Identification and Abundance Ratio Determinations.

Protein identification and quantitation is described in detail in a separate publication (53). Briefly, extracted proteins were combined in a 1:1 mass ratio to generate two samples: CO versus methanol and CO versus acetate. Then, these samples were separated by SDS/PAGE and each gel lane was cut into 10 bands of approximately similar density. Each band was in-gel-digested with trypsin, as described in ref. 54. Each sample was analyzed by liquid chromatography-tandem MS by using the Ultimate LC system (Dionex, Sunnyvale, CA) coupled to linear ion trap/Fourier-transform (LTQ-FT)-MS instrument (Thermo Electron, Waltham, MA), followed by separate sequential Sequest (55) searches for 15N and 14N peptides against the National Center for Biotechnology Information's database of M. acetivorans C2A (downloaded June 2005). Peptides with cross correlation values (Xcorr) >1.5 (1+), 2.0 (2+), and 2.5 (3+) and precursor mass within ±15 ppm of the theoretical mass initially were selected. Then, a protein list was composed, and proteins associated with probabilities >0.9 as calculated by ProteinProphet (56) were accepted as correct identifications. The protein abundance ratios were calculated based on areas of chromatographic peaks corresponding to individual peptides by using an in-house developed program, QN (53). For the most part, protein ratios were determined from at least two unique peptides. The average coefficient of variance of the observed protein abundance ratios was 9%, and approximately one-half of all proteins was quantitated with a coefficient of variance of <5%.

F420H2:CoB-S-S-CoM Oxidoreductase Assay.

Activity was monitored spectrophotometrically with a Beckman (Fullerton, CA) DU-7400 inside an anaerobic chamber (Coy Laboratory Products, Grass Lake MI) with a screw-cap cuvette. F420 was chemically reduced with NaBH4 as described in ref. 57. F420H2 oxidation was assayed under nitrogen in 0.3 ml of 50 mM Hepes, pH 7.5, containing 2 mM dithioerythritol and 20 μM F420H2. After adding 30–50 μg of cell extract, the cuvette was incubated for 5 min until a stable baseline was reached. The reaction was started by addition of 2 μl of CoB-S-S-CoM (final concentration, 90 μM) and followed spectrophotometrically at 420 nm. The reaction rate was calculated from an extinction coefficient of 42.5 mM−1 cm−1 for F420 at 420 nm.

Transcriptional Mapping.

Total RNA was isolated from CO-grown M. acetivorans cells and RT-PCR was performed as described in ref. 27. RNA ligation-mediated (RLM) RT-PCR was performed as described in ref. 58 except that RT-PCR was carried out with an Access RT-PCR kit (Promega, Madison, WI). Primers used are listed in Table 5, which is published as supporting information on the PNAS web site.

Quantitative RT-PCR.

Taqman assays were performed by using total RNA isolated from acetate-, methanol-, or CO-grown M. acetivorans C2A cells harvested at midexponential phase. Taqman primers and probes were designed with Primer Express 1.0 (Applied Biosystems, Foster City, CA). Primers were synthesized on the Bioautomation MerMade 12 (Plano, TX) at the Nucleic Acid Facility of the Pennsylvania State University. Probes were synthesized by Biosearch Technologies (Novato, CA) and were labeled with the reporter dye 6-carboxyfluorescein (6′-FAM) at the 5′ end and with the quencher dye Black Hole Quencher at the 3′ end. Primers and probes are summarized in Table 6, which is published as supporting information on the PNAS web site. cDNA was prepared by using the ABI High Capacity RT kit (Applied Biosystems) and was amplified and quantified on the ABI 7300 Real-Time PCR System (Applied Biosystems). Thermal cycling conditions were as follows: 2 min at 50°C, 10 min at 95°C, followed by 40 cycles of 15 s at 95°C and 1 min at 60°C. Relative abundances of each gene during growth on acetate, methanol, and CO were determined by using the ΔΔCT method (Applied Biosystems) using the 16s rRNA gene for normalization.

Supplementary Material

Supporting Tables:

Acknowledgments

We thank Lacy Daniels (Texas A & M University, Kingsville, TX) for generously supplying F420, Jan Keltjens (Radboud University Nijmegen, Nijmegen, The Netherlands) for providing CoB-S-S-CoM, and Deb Grove for assistance with Taqman assays. This work was supported by National Science Foundation Grant MCB-0110762 (to J.G.F.), a contract from Advanced Research International (to J.G.F.), and National Institutes of Health Grant GM15847 (to B.L.K.). This article is contribution number 885 from the Barnett Institute.

Abbreviations

THMPT
tetrahydromethanopterin
MF
methanofuran
CoB
coenzyme B
CoM
coenzyme M.

Footnotes

The authors declare no conflict of interest.

References

1. Conrad R. Microbiol Rev. 1996;60:609–640. [PMC free article] [PubMed]
2. Conrad R, Seiler W. Appl Environ Microbiol. 1980;40:437–445. [PMC free article] [PubMed]
3. Huber C, Wachtershauser G. Science. 1998;281:670–672. [PubMed]
4. Miyakawa S, Yamanashi H, Kobayashi K, Cleaves HJ, Miller SL. Proc Natl Acad Sci USA. 2002;99:14628–14631. [PMC free article] [PubMed]
5. Kasting JF. Science. 1993;259:920–926. [PubMed]
6. Russell MJ, Martin W. Trends Biochem Sci. 2004;29:358–363. [PubMed]
7. Ferry JG, House CH. Mol Biol Evol. 2006;23:1286–1292. [PubMed]
8. Meyer O, Frunzke K, Gadkari D, Jacobitz S, Hugendieck I, Kraut M. FEMS Microbiol Rev. 1990;87:253–260.
9. Dobbek H, Svetlitchnyi V, Gremer L, Huber R, Meyer O. Science. 2001;293:1281–1285. [PubMed]
10. Uffen RL. Enzyme Microb Technol. 1981;3:197–206.
11. Ludden PW, Roberts GP, Kerby RL, Spangler N, Fox J, Shelver D, He Y, Watt R. In: Microbial Growth on C1 Compounds. Lidstrom ME, Tabita FR, editors. Dordrecht, The Netherlands: Kluwer; 1996. pp. 183–190.
12. Wu M, Ren Q, Durkin AS, Daugherty SC, Brinkac LM, Dodson RJ, Madupu R, Sullivan SA, Kolonay JF, Nelson WC, et al. PLoS Genet. 2005;1:e65. [PMC free article] [PubMed]
13. Ljungdahl LG. In: Acetogenesis. Drake HL, editor. New York: Chapman & Hall; 1994. pp. 63–87.
14. Muller V, Gottschalk G. In: Acetogenesis. Drake HL, editor. New York: Chapman & Hall; 1994. pp. 127–156.
15. Parshina SN, Kijlstra S, Henstra AM, Sipma J, Plugge CM, Stams AJ. Appl Microbiol Biotechnol. 2005;68:390–396. [PubMed]
16. O'Brien JM, Wolkin RH, Moench TT, Morgan JB, Zeikus JG. J Bacteriol. 1984;158:373–375. [PMC free article] [PubMed]
17. Rother M, Metcalf WW. Proc Natl Acad Sci USA. 2004;101:16929–16934. [PMC free article] [PubMed]
18. Krzycki JA, Wolkin RH, Zeikus JG. J Bacteriol. 1982;149:247–254. [PMC free article] [PubMed]
19. Daniels L, Fuchs G, Thauer RK, Zeikus JG. J Bacteriol. 1977;132:118–126. [PMC free article] [PubMed]
20. Zeikus JG, Kerby R, Krzycki JA. Science. 1985;227:1167–1173. [PubMed]
21. Kluyver AJ, Schnellen CGTP. Arch Biochem. 1947;14:57–70. [PubMed]
22. Bott M, Thauer RK. Eur J Biochem. 1987;168:407–412. [PubMed]
23. Bott M, Thauer RK. Eur J Biochem. 1989;179:469–472. [PubMed]
24. Sowers KR, Baron SF, Ferry JG. Appl Environ Microbiol. 1984;47:971–978. [PMC free article] [PubMed]
25. Galagan JE, Nusbaum C, Roy A, Endrizzi MG, Macdonald P, FitzHugh W, Calvo S, Engels R, Smirnov S, Atnoor D, et al. Genome Res. 2002;12:532–542. [PMC free article] [PubMed]
26. Ferry JG. In: Biochemistry and Physiology of Anaerobic Bacteria. Ljungdahl LG, Adams MW, Barton LL, Ferry JG, Johnson MK, editors. New York: Springer; 2003. pp. 143–156.
27. Li Q, Li L, Rejtar T, Lessner DJ, Karger BL, Ferry JG. J Bacteriol. 2006;188:702–710. [PMC free article] [PubMed]
28. Li Q, Li L, Rejtar T, Karger BL, Ferry JG. J Proteome Res. 2005;4:129–136. [PubMed]
29. Li Q, Li L, Rejtar T, Karger BL, Ferry JG. J Proteome Res. 2005;4:112–128. [PubMed]
30. Gottschalk G, Thauer RK. Biochim Biophys Acta. 2001;1505:28–36. [PubMed]
31. Welander PV, Metcalf WW. Proc Natl Acad Sci USA. 2005;102:10664–10669. [PMC free article] [PubMed]
32. Kerby RL, Hong SS, Ensign SA, Coppoc LJ, Ludden PW, Roberts GP. J Bacteriol. 1992;174:5284–5294. [PMC free article] [PubMed]
33. Ensign SA, Ludden PW. J Biol Chem. 1991;266:18395–18403. [PubMed]
34. Terlesky KC, Nelson MJK, Ferry JG. J Bacteriol. 1986;168:1053–1058. [PMC free article] [PubMed]
35. Maupin-Furlow JA, Ferry JG. J Bacteriol. 1996;178:6849–6856. [PMC free article] [PubMed]
36. Fischer R, Thauer RK. FEBS Lett. 1990;269:368–372. [PubMed]
37. Terlesky KC, Ferry JG. J Biol Chem. 1988;263:4075–4079. [PubMed]
38. Stojanowic A, Hedderich R. FEMS Microbiol Lett. 2004;235:163–167. [PubMed]
39. Meuer J, Kuettner HC, Zhang JK, Hedderich R, Metcalf WW. Proc Natl Acad Sci USA. 2002;99:5632–5637. [PMC free article] [PubMed]
40. Deppenmeier U. J Bionenerg Biomembr. 2004;36:55–64. [PubMed]
41. Simianu M, Murakami E, Brewer JM, Ragsdale SW. Biochemistry. 1998;37:10027–10039. [PubMed]
42. Heiden S, Hedderich R, Setzke E, Thauer RK. Eur J Biochem. 1993;213:529–535. [PubMed]
43. Stojanowic A, Mander GJ, Duin EC, Hedderich R. Arch Microbiol. 2003;180:194–203. [PubMed]
44. Abbanat DR, Ferry JG. J Bacteriol. 1990;172:7145–7150. [PMC free article] [PubMed]
45. Deppenmeier U, Johann A, Hartsch T, Merkl R, Schmitz RA, Martinez-Arias R, Henne A, Wiezer A, Baumer S, Jacobi C, et al. J Mol Microbiol Biotechnol. 2002;4:453–461. [PubMed]
46. Deppenmeier U. Prog Nucleic Acid Res Mol Biol. 2002;71:223–283. [PubMed]
47. Hedderich R, Berkessel A, Thauer RK. Eur J Biochem. 1990;193:255–261. [PubMed]
48. Hedderich R. J Bionenerg Biomembr. 2004;36:65–75. [PubMed]
49. Zinder S. In: Methanogenesis. Ferry JG, editor. New York: Chapman & Hall; 1993. pp. 128–206.
50. Sowers KR, Boone JE, Gunsalus RP. Appl Environ Microbiol. 1993;59:3832–3839. [PMC free article] [PubMed]
51. Ferry JG, Wolfe RS. Arch Microbiol. 1976;107:33–40. [PubMed]
52. Grant WM. Anal Chem. 1948;20:267–269.
53. Andreev VP, Li L, Rejtar T, Li Q, Ferry JG, Karger BL. J Proteome Res. 2006;5:2039–2045. [PubMed]
54. Shevchenko A, Wilm M, Vorm O, Mann M. Anal Chem. 1996;68:850–858. [PubMed]
55. Eng JK, McCormack AL, Yates JR. J Am Soc Mass Spectrom. 1994;5:976–989. [PubMed]
56. Nesvizhskii AI, Keller A, Kolker E, Aebersold R. Anal Chem. 2003;75:4646–4658. [PubMed]
57. Deppenmeier U, Blaut M, Mahlmann A, Gottschalk G. FEBS Lett. 1990;261:199–203.
58. Bensing BA, Meyer BJ, Dunny GM. Proc Natl Acad Sci USA. 1996;93:7794–7799. [PMC free article] [PubMed]

Articles from Proceedings of the National Academy of Sciences of the United States of America are provided here courtesy of National Academy of Sciences
PubReader format: click here to try

Formats:

Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...

Links

Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...