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Keywords:

  • methanogenesis;
  • tungstoenzymes;
  • interspecies hydrogen transfer;
  • formate;
  • syntrophy

Abstract

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

Two formate dehydrogenases (CO2-reductases) (FDH-1 and FDH-2) were isolated from the syntrophic propionate-oxidizing bacterium Syntrophobacter fumaroxidans. Both enzymes were produced in axenic fumarate-grown cells as well as in cells which were grown syntrophically on propionate with Methanospirillum hungatei as the H2 and formate scavenger. The purified enzymes exhibited extremely high formate-oxidation and CO2-reduction rates, and low Km values for formate. For the enzyme designated FDH-1, a specific formate oxidation rate of 700 U·mg−1 and a Km for formate of 0.04 mm were measured when benzyl viologen was used as an artificial electron acceptor. The enzyme designated FDH-2 oxidized formate with a specific activity of 2700 U·mg−1 and a Km of 0.01 mm for formate with benzyl viologen as electron acceptor. The specific CO2-reduction (to formate) rates measured for FDH-1 and FDH-2, using dithionite-reduced methyl viologen as the electron donor, were 900 U·mg−1 and 89 U·mg−1, respectively. From gel filtration and polyacrylamide gel electrophoresis it was concluded that FDH-1 is composed of three subunits (89 ± 3, 56 ± 2 and 19 ± 1 kDa) and has a native molecular mass of approximately 350 kDa. FDH-2 appeared to be a heterodimer composed of a 92 ± 3 kDa and a 33 ± 2 kDa subunit. Both enzymes contained tungsten and selenium, while molybdenum was not detected. EPR spectroscopy suggested that FDH-1 contains at least four [2Fe-2S] clusters per molecule and additionally paramagnetically coupled [4Fe-4S] clusters. FDH-2 contains at least two [4Fe-4S] clusters per molecule. As both enzymes are produced under all growth conditions tested, but with differences in levels, expression may depend on unknown parameters.

Abbreviations
FDH

formate dehydrogenases (CO2-reductases)

During methanogenic decomposition, only a small amount of energy becomes available for the microorganisms involved. Some of the reactions are even endergonic under thermodynamic standard conditions (298 K, pH 7, concentrations of 1 m and partial pressures of 105 Pa for gases). In particular, some reduced organic products of fermenting organisms such as acetate, propionate, butyrate and benzoate require extremely low product concentrations in order to make oxidation of these compounds energetically favorable. Nevertheless, some organisms are able to gain metabolic energy by oxidizing these compounds, provided the products are removed efficiently. This type of symbiotic association, in which the partners depend on each other to perform the metabolic activity observed, was termed syntrophy. The partner organisms of syntrophic acetogenic bacteria in methanogenic environments are hydrogenotrophic methanogenic archaea [1,2]. Because H2 is produced by the acetogens and consumed by the methanogens and the concentration of this compound in particular has to be extremely low, this process is often referred to as ‘interspecies H2-transfer’. Formate, however, is also considered to be an important mediator of reducing equivalents in syntrophic cultures [3–8]. As the concentrations of both H2 and formate in these cultures are very low, it is extremely difficult to determine which species is preferred. If formate is transferred, the acetogens should possess formate dehydrogenases (FDHs; CO2-reductases), which are active in the direction of CO2-reduction. The energetically unfavorable reactions are thought to be driven by reversed electron transport during syntrophic growth. Because a membrane-system would make it possible to shift electrons to a lower redox potential, suitable to couple succinate oxidation to proton or bicarbonate reduction [2,9], these enzymes are probably periplasmic.

FDH catalyzes the oxidation of formate to CO2; the reverse reaction, the reduction of CO2 to formate, has only been demonstrated for a few FDHs. FDH has been purified from several sources including eukaryotes, archaea and bacteria. In general, the FDHs from aerobic organisms reduce NAD+, have a high Km for formate and are insensitive to O2[10]. Metals and cofactors (which are all flavins) were found in only a few of these enzymes [10–13]. In contrast, all the FDHs isolated from anaerobic bacteria and archaea contain complex redox centers, they are extremely sensitive to O2 and most of them do not reduce NAD+. Apart from multiple iron-sulfur clusters, FDHs from anaerobes contain molybdenum in their active sites, usually in combination with selenium [14]. However, the NADP-reducing FDH of Moorella thermoacetica (previously known as Clostridium thermoaceticum), contains tungsten [15], and recently two other W-containing FDHs were isolated [16,17]. One of these enzymes was isolated from the sulfate reducer Desulfovibrio gigas, while the FDHs of Desulfovibrio vulgaris Hildenborough and Desulfovibrio desulfuricans are molybdoenzymes [18,19]. Almendra et al. (1999) hypothesized that extant Desulfovibrio species preferably produce Mo-containing proteins, while their predecessors in evolution utilized both tungsten and molybdenum [16].

Syntrophobacter fumaroxidans is a syntrophic propionate-oxidizer isolated from anaerobic granular sludge [20]. It grows in suspended cocultures with methanogens that utilize both H2 and formate [6]. Phylogenetic analysis revealed that S. fumaroxidans as well as other Syntrophobacter species are closely related to sulfate-reducing bacteria [20–22]. These organisms are indeed able to couple propionate oxidation to sulfate reduction, though at a much lower rate than other sulfate reducers [23–25]. S. fumaroxidans possesses at least two formate dehydrogenases, of which the expression levels appeared to be much higher during syntrophic growth than in cells that were cultured axenically [8,26]. To date there are no reports, however, on the isolation and properties of an FDH required for syntrophic growth. We report here the purification and properties of two distinct, W-containing FDHs from S. fumaroxidans. These enzymes exhibit extremely high CO2-reduction and formate-oxidation rates as compared with the FDHs described for other anaerobic bacteria.

Organism and growth conditions

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

S. fumaroxidans (DSM 10017) and cocultures of S. fumaroxidans with Methanospirillum hungatei JF1 (DSM 864) were grown in bicarbonate-buffered medium with the following composition: 3 mm Na2HPO4, 3 mm KH2PO4, 5.6 mm NH4Cl, 0.75 mm CaCl2, 0.5 mm MgCl2, 5 mm NaCl, 50 mm NaHCO3, 1 mm Na2S, 7.5 µm FeCl2, 1 µm H3BO3, 0.5 µm ZnCl2, 0.1 µm CuCl2, 0.5 µm MnCl2, 0.5 µm CoCl2, 0.1 µm NiCl2, 0.1 µm Na2SeO3, 0.1 µm Na2WO4, 0.1 µm Na2MoO4, 0.5 mg·L−1 EDTA, vitamins (µg·L−1); 0.02 biotin, 0.2 nicotinic acid, 0.5 pyridoxine, 0.1 riboflavin, 0.2 thiamin, 0.1 cyanocobalamin, 0.1 p-aminobenzoic acid, 0.1 pantothenic acid, 0.1 lipoic acid and 0.1 folic acid. For large-scale cultivation the cultures were grown in 25-L carboys containing 20 L of medium with 30 mm fumarate as the substrate for S. fumaroxidans and 30 mm propionate for methanogenic cocultures. Per 20 L of culture medium, the axenic culture produced approximately 20 g of wet cell-pellet on fumarate, while in the coculture the yield was approximately 4 g of cells on propionate (ratio of S. fumaroxidans/M. hungatei≈ 1 : 1). The methanogens were pregrown on 10 mm of sodium formate, prior to inoculation of syntrophic coculture. Cells were harvested anoxically by continuous-flow centrifugation and stored at −20 °C after resuspending the cell pellets in 50 mm Tris/HCl, pH 8.

Enzyme purification

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

All purification procedures were performed in an anaerobic glove box with N2/H2 (96 : 4, v/v) as the gas phase. The oxygen concentration was kept below 2 p.p.m. by circulating the gas phase over a platinum catalyst column. Thawed cells were broken in an Aminco French pressure cell at 100 MPa. After treatment with deoxyribonuclease the extracts were centrifuged at 16 000 g for 10 min to remove cell debris. The enzymes were purified from the soluble fraction of the cells, which was obtained by ultracentrifugation of the crude extracts at 140 000 g and 4 °C for 1 h. All buffers used for purification were degassed and filtered through a 45-µm filter, and after equilibration in the anaerobic glove box for at least 24 h, supplied with 150 µm sodium dithionite prior to use. The soluble fraction was loaded onto a Q-sepharose fast-flow column (2 × 10 cm) equilibrated with 50 mm Tris/HCl pH 8 (buffer A). A 320-mL linear gradient of 0–0.6 m NaCl in buffer A was applied followed by an 80-mL linear gradient of 0.6–1 m NaCl. Fractions containing FDH, which eluted between 0.16 and 0.30 m NaCl, were pooled and applied to a ceramic hydroxyapatite column (Biogel-CHT5, 1 × 6.4 cm, Bio-Rad) equilibrated with 0.01 mm Na/Pi, pH 7, at a flow rate of 1 mL·min−1. The FDH that did not bind to this column was designated FDH-1. The other FDH, which eluted at 0.32 m phosphate in a 75-mL linear gradient of 0.01–0.45 m Na/phosphate, was designated FDH-2. FDH-1 was concentrated and desalted in a Filtron 30-kDa ultrafiltration cell. For further purification the enzyme was applied to a Mono-Q column equilibrated with buffer A, and eluted with a 20-mL linear gradient of 0–0.35 m NaCl at a flow rate of 1 mL·min−1 in the same buffer. FDH-1 eluted at 0.16 m NaCl and was concentrated in an Amicon ultrafiltration cell with a PM30 filter and applied to a Superdex 200 pg (1 × 30 cm, preparative grade, Amersham Pharmacia) column equilibrated with buffer A containing 0.15 m NaCl. A flow rate of 0.5 mL·min−1 was applied to elute the protein. For purification of FDH-2 we repeated the first two steps (Q-sepharose and Biogel-CHT), starting with fresh material. For further purification of FDH-2 the same principles and conditions as described for FDH-1 were applied. FDH-2 eluted at 0.24 m Cl from the Mono-Q and as a single peak from the Superdex 200 pg. Purified enzymes were either stored under N2 at 0 °C (for all characterization procedures carried out within the next 24 h), or frozen in liquid N2 and then stored for longer periods at −70 °C. For purification of FDH-1 and FDH-2 from cells cocultured with M. hungatei, the purification procedures applied were similar to those for fumarate-grown cells.

Quantification of the formate dehydrogenase levels

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

A ceramic hydroxyapatite column was used to quantify the levels of FDH-1 and FDH-2 in S. fumaroxidans cells which were grown syntrophically, as described previously for axenically cultured S. fumaroxidans cells [26]. Syntrophobacter cells were separated from M. hungatei by Percoll gradient centrifugation as described previously [8]. The S. fumaroxidans cells (0.1 g) were resuspended in 1.5 mL 50 mm Tris/HCl, pH 8, and disrupted by sonication. The soluble fraction after centrifugation at 140 000 g (4 °C for 1 h) was loaded directly onto the ceramic hydroxyapatite column described in the previous section, using similar conditions. In this case both FDHs bound to the column (FDH-1 did not bind to this column when separation on this particular medium was preceded by anion exchange chromatography). Proteins were eluted from the column at a flow rate of 1 mL·min−1 in a 50-mL linear gradient of 0.01–0.25 m sodium phosphate, followed by a 20-mL linear gradient of 0.25–0.5 m sodium phosphate. The proteins that eluted from the column were collected in 3.5-mL fractions and subsequently analyzed for formate dehydrogenase activity. FDH-1 and FDH-2 eluted in separate peaks at 210 mm phosphate and 300 mm phosphate, respectively [26].

Enzyme assays and kinetic analyses

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

Enzyme activities were measured at 37 °C in N2-flushed cuvettes closed with butyl-rubber stoppers. One unit is defined as the amount of enzyme catalyzing the conversion of 1 µmol of substrate per min. During purification FDH activity was followed by measuring the reduction of 1 mm benzyl viologen (BV2+) in 50 mm Tris/HCl, pH 8, at 578 nm (BV+, ε = 8.65 mm−1·cm−1). Before addition of the enzyme the assay buffer was slightly prereduced with sodium dithionite to obtain an absorption of 0.2–0.5. The reaction was initiated by addition of 10 mm sodium formate. Other artificial electron acceptors tested in the same buffer using a similar procedure as for BV were; 1 mm methyl viologen (MV+, ε = 9.7 mm−1·cm−1 at 578 nm); 3 mm potassium ferricyanide; 0.5 mm anthraquinone-2,6-disulfonic acid; 0.2 mm 2,6-dichlorophenolindophenol; 0.5 mm flavin mono nucleotide; 41 mm FAD; 1 mm NAD and 0.125 mg·mL−1 methanogenic cofactor F420. The CO2-reductase activity was measured by following the oxidation of dithionite-reduced methyl viologen in 100 mm sodium phosphate, pH 7.3, at 578 nm. The reaction was started by the addition of 10 mm sodium bicarbonate. Kinetic parameters were calculated from Lineweaver–Burk plots obtained from activity measurements of the purified enzymes at varying substrate or electron acceptor concentrations. Protein was determined according to Bradford [27] with bovine serum albumin as a standard.

Molecular composition of the purified formate dehydrogenases

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

The molecular masses of the purified FDHs were estimated from gel filtration and SDS/PAGE according to Laemmli [28]. A combination of high and low molecular mass markers used as a reference for gel filtration consisted of (size in kDa); blue dextran, 2000; thyroglobulin, 669; ferritin, 440; catalase, 232; aldolase 158; bovine serum albumin, 67; ovalbumin, 43; chymotrypsinogen, 25 and ribonuclease 13.7. A low molecular mass marker (Bio-Rad) was used as a reference for SDS gel electrophoresis. The metal content of the purified enzymes was measured by inductively coupled plasma mass spectrometry (Elan 6000, Perkin-Elmer) [29]. For those preparations the protein content was determined with a microbiuret method [30].

Staining of formate dehydrogenase in polyacrylamide gels

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

Staining of FDH activity after nondenaturating PAGE was performed at room temperature under a nitrogen atmosphere. The gel was incubated in Tris/HCl pH 8, containing 1 mm methyl viologen which was slightly prereduced with sodium dithionite. Staining was started by addition of 1 mm triphenyltetrazolium chloride and subsequently 10 mm sodium formate. Heme was stained with tetramethylbenzidine according to Trumpower and Katki [31] after separation of the FDH subunits in an LDS (lithium dodecyl sulphate) polyacrylamide gel.

N-terminal amino acid sequencing

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

Purified proteins were transferred from SDS/polyacrylamide gels (12% for large subunits and 15% for small subunits) to a polyvinylidene difluoride membrane in a Bio-Rad Mini Trans-Blot module. Transfer was carried out at 200 mA for 3 h in 10 mm CAPS buffer pH 11 containing 2.5 m methanol. Immobilized proteins were stained with Ponceau S. The N-terminal sequences of the FDH subunits were determined by Edman-degradation as described previously [32].

Purification of the Syntrophobacter fumaroxidans formate dehydrogenases

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

Purification of FDH activity from the soluble fraction of Syntrophobacter fumaroxidans cells grown on fumarate yielded two distinct enzymes, FDH-1 and FDH-2. Strictly anoxic conditions and the addition of dithionite as a reductant to the buffers were required to recover FDH activity. Other reducing agents such as dithiothreitol, 2-mercaptoethanol, sulfide or cysteine had an insufficient stabilizing effect on the enzyme activity during purification. In addition, oxygen-inactivated enzyme preparations could not be reactivated with any of the reductants mentioned above. In order to fully activate the enzymes an initial reduction of the viologen dyes was required, otherwise no or a much lower activity was measured. Furthermore, maximal rates were only measured after an incubation period (without substrate) of usually 2–5 min until a stable baseline was obtained. Storage of partially purified FDH under N2 at 0 °C, either with or without reducing agents, could not overcome rapid inactivation of the enzymes in time. Therefore, though the use of the same purification steps for each enzyme would suggest simultaneous purification of both enzymes, we started with fresh material for the purification of each individual enzyme.

FDH-2 of S. fumaroxidans was purified 79-fold from fumarate-grown cells to a specific activity of 2700 U·mg−1(Table 1). Purification of this enzyme from cells cocultured with M. hungatei was less successful, probably due to the presence of the archaeal proteins. With respect to purity, however, for FDH-1 the best results were obtained when purified from cocultured cells (Table 1). When this enzyme was purified from fumarate-grown cells, two contaminating bands were still visible in the SDS/polyacrylamide gels after electrophoresis (Fig. 1A). Though active FDH could be recovered from mild hydrophobic interaction chromatography media, such an additional step was not successful to remove these contaminating protein bands. The highest specific activity measured for this enzyme was 700 U·mg−1 when purified 13-fold from cells cocultured with M. hungatei. The low recoveries of 2 and 6% for FDH-1 and FDH-2, respectively, indicate that a large amount of activity was lost upon purification. Activity-staining of the isolated enzymes after native polyacrylamide gel electrophoresis showed that the bands observed with Coomassie staining also showed FDH activity (Fig. 1B). Heme was not detected in the subunits of the isolated enzymes, nor did we observe bands typical for heme in the absorption spectra of the isolated enzymes.

Table 1. Purification of the formate dehydrogenases of syntrophobacter fumaroxidans. Purification of FDH-1 was from S. fumaroxidans cells grown on propionate in coculture with Methanospirillum hungatei. Purification was started with 6 g coculture cells. Purification of FDH-2 was from 12 g of S. fumaroxidans cells grown on fumarate.
FractionFDH-1FDH-2
Total activity (U)Specific activity (U·mg−1)Total activity (U)Specific activity (U·mg−1)
  • a

    These values represent the combined activities of the S. fumaroxidans and the M. hungatei formate dehydrogenases.

140 000 g25 763a 55a1330034
Q-sepharose11 714691087277
Biogel-CHT53240512340813
Mono-Q150519725341274
Superdex 2005046988502659
image

Figure 1. Polyacrylamide gel electrophoresis of the S. fumaroxidans formate dehydrogenases. (A) Enzymes separated under SDS-denaturing conditions. 1, Low molecular mass marker (the arrow indicates a 50 kDa protein); 2, FDH-1 purified from fumarate-grown cells; 3, FDH-1 purified from propionate-grown cells (cocultured with M. hungatei); 4, FDH-2 purified from fumarate-grown cells. (B) Electrophoresis under nondenaturing conditions followed by activity staining. 1 and 3, FDH-1; 2 and 4, FDH-2. Protein stained with Coomassie (1 and 2), or for activity with triphenyltetrazoliumchloride (3 and 4).

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Quantification of the formate dehydrogenase levels

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

The specific activity of FDH in the soluble fraction of cocultured S. fumaroxidans cells, which were separated by Percoll gradient centrifugation, was 92 U·mg−1. In axenic cultures these levels were 10-fold lower [26]. By separating the FDHs of syntrophically grown S. fumaroxidans cells on a ceramic hydroxyapatite column, it was estimated that approximately 75% of the total FDH activity corresponded with FDH-1 and the remaining 25% with FDH-2. This ratio was comparable to the ratio of both FDHs in S. fumaroxidans cells grown on other substrates in the absence of methanogens [26].

Characterization of S. fumaroxidans formate dehydrogenases

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

Methyl and benzyl viologen served as electron acceptors for both isolated S. fumaroxidans enzymes (Table 2). 2,6-Dichloroindophenol (9.6%), anthraquinone-2,6-disulfonate (not quantified), ferricyanide (36%), FMN (1.3%) and FAD (8.1%) could also be used to measure FDH in the soluble extract (the percentages of activity measured as compared to the activity measured with benzyl viologen are presented between brackets), but not for preparations of the (partially) purified enzymes. Both enzymes, and enriched fractions of the enzymes in particular, required a reactivation time of up to 10 min in order to measure the maximum formate oxidation rates. Reactivation was always accompanied by an increase of reduced dye. NAD(P) and F420 did not support the oxidation of formate, which was also confirmed by measurements with soluble extracts. Only methyl viologen supported the reduction of CO2, which was catalyzed by both FDHs, with dithionite as electron donor (Table 2). The CO2 concentration in the assay buffer under these conditions was approximately 1 mm, assuming equilibrium between [CO2] and [HCO3].

Table 2. Formate oxidation and CO2-reduction rates of the isolated formate dehydrogenases. All reactions were measured spectrophotometrically at 37 °C and 578 nm; formate oxidation rates with 1 mm methyl viologen and 10 mm formate, CO2 reduction rates with 0.5 mm dithionite reduced methyl viologen and 10 mm sodium bicarbonate.
Donor/acceptorpHFDH-1 (U·mg−1)FDH-2 (U·mg−1)
Formate/BV2+87002700
Formate/MV2+8930510
Formate/MV2+7.354468
MV+/HCO3 (CO2)7.390089

Molecular mass and composition of the isolated enzymes were estimated using gel filtration in combination with SDS/PAGE. FDH-2 consisted of two subunits of 33 ± 2 kDa and 92 ± 3 kDa (Fig. 1A), while a molecular mass of 100 kDa estimated from gel filtration suggests a molecular mass of 125 ± 5 kDa for the native enzyme. The subunit sizes of FDH-1 are 89 ± 3, 56 ± 2 and 19 ± 1 kDa (Fig. 1A), whereas a molecular mass around 350 kDa was estimated for the native enzyme. A possible composition of the native enzyme is a α2β2γ2 structure with a molecular mass of 328 kDa. Metal analysis revealed the presence of tungsten and selenium in both enzymes, while molybdenum was not detectable (Table 3). The amount of W per molecule was less than 1 for both enzyme preparations of FDH-1 tested, but the proteins of these samples (purified from fumarate-grown cells) were not completely pure (Fig. 1A). The amount of selenium was 1.5–2.5 times as much as the amount of W present in these samples, suggesting a 1 : 2 ratio of W and Se in FDH-1. The amount of iron in this enzyme is uncertain as there is a large difference between the two preparations of FDH-1 that were analyzed. The amounts of W and Se detected in FDH-2 seemed to be consistent with one W and one Se per molecule, and this enzyme contained 16–18 iron atoms per heterodimer (Table 3). Km values for formate determined at pH 8 with 5 mm benzyl viologen as electron acceptor were 0.04 mm for FDH-1 and 0.01 mm for FDH-2. Due to the reactivation time required for the purified enzymes, we were not able to determine Km values for CO2 (or bicarbonate).

Table 3. Metal content of the isolated formate dehydrogenases of Syntrophobacter fumaroxidans (mol·protomer−1, assuming that the molecular masses calculated for FDH-1 and FDH-2 are 350 kDa and 125 kDa, respectively) as determined with inductively coupled mass spectrometry. The amount of protein analyzed on the ICP-MS is indicated in the second column.
Purified enzymeProtein (mg)FeSeWMo
  • a

     The samples of formate dehydrogenase 1 used for metal analysis contained small amounts of contaminating protein.

FDH-1a0.33480.90.60.0
FDH-1a1.05371.20.50.0
FDH-22.56160.70.60.0
FDH-20.69181.31.00.0

N-terminal amino acid sequences were obtained for all five subunits of the S. fumaroxidans FDHs (Table 4). Sequence similarity searches and alignments were performed using the blast program and clustalw. The small subunit of FDH-2 revealed highest similarity to the 29 kDa α-subunit from the periplasmic Desulfovibrio vulgaris Hildenborough FDH. Remarkably the large subunit of FDH-1 showed highest similarity to the smaller subunit of one of the Eubacterium acidaminophilum FDHs (Table 4). The similarities of the other N-terminal sequences to those of related sequences were much less significant.

Table 4. N-terminal sequences of the subunits of the S. fumaroxidans formate dehydrogenases. The amino acids marked ‘X’ could not be determined and those indicated by small letters are not unambiguous. Related sequences were obtained from the NCBI GenBank database and aligned using the clustal w program. Accession numbers (NCBI); E. acidaminophilum fdhB-II; CAC39238, D. vulgaris fdh-β; AAB35992. Conserved amino acids are shown in bold.
SubunitSequence
S. fumaroxidans FDH-1  α-subunit 89 kDaMDNNIXTLKVNGQSVKXHK
E. acidaminophilum fdhB-IIMVTLTINGQSVSVSR
S. fumaroxidans FDH-1  β-subunit 59 kDaMQPQASilKvPAXEXgVL
S. fumaroxidans FDH-1  γ-subunit 19 kDaMXXXLqQIFTtXDDXXXALIPV
S. fumaroxidans FDH-2  α-subunit 92 kDaYTXELXTKDAKETPXIDXYKAk
S. fumaroxidans FDH-2  β-subunit 32 kDaAGKSFFIDTTGXTAkXGDQ
D. vulgaris Hildenborough fdh  β-subunitKAFLIDTTRXTAXRGXQ

EPR spectroscopy

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

Low temperature EPR of the isolated FDH-1 was dominated by an axial S = 1/2 signal with g-values 2.01, 1.94 and 1.94, indicative of a [4Fe-4S]+ or [2Fe-2S]+ cluster (Fig. 2, Table 5). Furthermore the S = 1/2 signal was flanked with features suggesting the presence of one or more other FeS clusters. At 52 K only the axial S = 1/2 signal was found (data not shown). This axial S = 1/2 signal was attributed to a [2Fe-2S]+ cluster, as EPR signals from a [4Fe-4S]+ cluster broaden severely above 40 K. Reduction with sodium formate resulted in an additional rhombic S = 1/2 EPR signal, with g-values 2.00, 1.95 and 1.92, which was also not broadened at 54 K (Fig. 3). At low temperatures additional features in the g = 2 region appeared, which indicated the presence of [4Fe-4S]+ clusters. However, Almendra et al. [16] interpreted similar signals from a tungsten-containing FDH from D. gigas as originating from two [4Fe-4S] clusters. The EPR signals found for this FDH are remarkably similar to those found for the tungsten-containing FDH from Moorella thermoacetica[36]. These authors interpreted the EPR data with two [2Fe-2S] clusters and two [4Fe-4S] clusters. The EPR spectra of FDH-1 at low temperature, however, was more complicated most likely due to coupling to a fast relaxing paramagnet such as a [4Fe-4S] cluster. Double integration of the EPR signals at 54 K with reference to an external copper standard affords a spin quantitation of 2.3 spins per αβγ unit for [2Fe-2S]I and [2Fe-2S]II together. The total spectrum at low temperature doubly integrated to 8.6 spins per αβγ unit, indicating FDH-1 may contain additionally six [4Fe-4S] clusters.

image

Figure 2. EPR spectra of S. fumaroxidans FDH-1. Trace A, as isolated enzyme at 16.5 K. Trace B, reduced with sodium formate at 22 K. Trace C, formate-reduced enzyme at 54 K. Microwave frequency, 9.22 GHz; microwave power, 7.96 mW; modulation frequency, 100 kHz; modulation amplitude, 1.0 mT.

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Table 5. Simulation parameters of the EPR signals from S. fumaroxidans FDH-1 and FDH-2. The simulations assume S = 1/2 and are based on 100 × 100 molecular orientations. The line shape is assumed to be a symmetrical Gaussian in frequency space; the line width is described as a g-stain tensor colinear with the g-tensor (50).
 g-ValueLine width
gzgygxΔzΔyΔx
FDH-1
 [2Fe-2S]I2.0131.9371.9370.00500.00560.0056
 [2Fe-2S]II2.0031.9531.9220.00450.00700.0066
FDH-2
 [4Fe-4S]2.0441.9411.8860.0070.0060.007
 Radical2.0012.0012.0010.0060.0060.006
image

Figure 3. EPR spectra of formate-reduced S. fumaroxidans FDH-1. Trace A, experimental spectrum. Trace B, simulation of [2Fe-2S]I + [2Fe-2S]II. Trace C, simulation of [2Fe-2S]I. Trace D, simulation of [2Fe-2S]II. Microwave frequency, 9.22 GHz; microwave power, 12.6 mW; modulation frequency, 100 kHz; modulation amplitude, 0.63 mT; temperature 54 K.

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Low temperature EPR of an isolated FDH-2 showed a rhombic S = 1/2 EPR signal, with g-values 2.04, 1.94 and 1.89 (Fig. 4), which rapidly broadened at temperatures above 40 K (data not shown). Both the g-values and the saturation characteristics were indicative of a [4Fe-4S]+ cluster. Furthermore a slowly relaxing isotropic signal with g = 2.00 was found, which clearly originated from a radical. Double integration of the EPR signals resulted in 2.1 spins per molecule for the [4Fe-4S]+ cluster and 0.1 spins per molecule for the radical. More detailed measurement of the radical under nonsaturation conditions did not show any (partially) resolved hyperfine splitting. The peak-to-peak width of the signal is 15 G, which seems consistent with a flavin radical. However, chemical analysis did not show the presence of flavin in this enzyme. Reduction with sodium formate resulted in only small changes of the EPR spectrum: the appearance of an additional weak S = 1/2 signal with gy = 1.88 and gx = 1.85 was observed. This S = 1/2 signal does not broaden at 60 K (data not shown), therefore it may represent W5+ which is a slowly relaxing species.

image

Figure 4. EPR spectra of S. fumaroxidans FDH-2. Trace A, as isolated. Trace B, reduced with sodium formate. Trace C, simulation. Microwave frequency, 9.22 GHz; microwave power, 3.17 mW; modulation frequency, 100 kHz; modulation amplitude, 1.0 mT; temperature, 22 K.

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Discussion

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References

The syntrophic propionate-oxidizing bacterium Syntrophobacter fumaroxidans possesses two distinct W-containing FDHs with very high specific activities. Both enzymes were present in fumarate-grown cells as well as in cells grown syntrophically on propionate with Methanospirillum hungatei as the H2 and formate scavenger. The assay used to follow the viologen-dependent oxidation of formate was only successful if the dyes were already partially reduced prior to the addition of formate, and in addition, a reactivation time of up to 10 min was required to measure the maximum activity. Apparently both enzymes required reducing conditions as described for the tungsten-containing acetylene hydratase from Pelobacter acetylenicus[37]. Such reducing conditions in the assay were also required to prevent a lag phase in the activity of the W-containing aldehyde dehydrogenase from Desulfovibrio gigas. Similar reducing conditions were also required to maintain the enzyme stability during purification: a concentration of 100 µm dithionite in the buffers was sufficient to obtain reasonable (> 50%) recoveries per purification step. Unlike the W-containing FDH from D. gigas[16,38], strong reducing conditions were also required to purify the W-containing FDHs from M. thermoacetica[15] and E. acidaminophilum[17].

Although syntrophic organisms have been shown to possess FDH activity [8,9,39,40] the only biochemical evidence for formate transfer was obtained only recently [8]. It has been calculated that due to the low diffusion rate of H2, formate is a more obvious electron carrier in suspended cocultures [5]. Therefore, we believe that syntrophic propionate oxidizers are able to reduce both protons and bicarbonate during syntrophic growth. The syntrophic partner organism and the interbacterial distance may determine which mechanism prevails under the specified conditions. The presence of FDHs in S. fumaroxidans, which are very active in the direction of CO2-reduction and of which the levels are highest during syntrophic growth in suspended cocultures [8], supports this theory. Here we demonstrated that the levels of both FDHs are higher during syntrophic growth. We speculate that the organism needs two FDHs to grow on propionate syntrophically [26]. One of the enzymes is required to dispose of reducing equivalents as formate, and is therefore probably located in the periplasm. The other FDH may be required to fix CO2 via the reverse acetyl-CoA cleavage pathway. In S. fumaroxidans the presence of this pathway was demonstrated in cells grown on fumarate [41]. During syntrophic growth on propionate, the organism uses an acetyl-CoA/propionate HS-CoA transferase to activate propionate, while other possible enzymes to activate propionate directly have not been detected. Propionate is thus converted stoichiometrically to acetate, and therefore the Wood pathway is thought to have an anaplerotic function in this organism during growth on propionate [41]. Localization experiments with membrane-impermeable viologen dyes suggested that neither of the two enzymes is periplasmic [26]. In the present study we tried to wash periplasmic enzymes from intact cells as described for Desulfovibrio vulgaris FDH [18], but S. fumaroxidans periplasmic enzymes were not released by using this procedure. Attempts to prepare spheroplasts of S. fumaroxidans cells were also not successful as the procedure tested [42] resulted in aggregation of the cells (data not shown). Unfortunately, our attempts to identify the genes encoding the S. fumaroxidans FDHs have also not been successful so far. However, with the N-terminus of the FDH-2 large subunit, we managed to isolate a fragment of the FDH-2 encoding gene (Fritsche et al. unpublished results). A twin-arginine motif upstream of the N-terminal sequence, which was determined by Edman degradation, suggests that FDH-2 is translocated to the periplasm upon maturation [43]. We therefore expect that FDH-2 will be involved in the disposal of reducing equivalents and that FDH-1 will be active in the reductive CO2-fixation pathway. Further experiments are planned to address these issues.

Both S. fumaroxidans FDHs appear to be tungsten-selenium enzymes. Although both tungsten and molybdenum were present in the growth medium, only tungsten was detected in the purified enzymes. The metal content of FDH-1 may be underestimated because the preparations used for analysis of the metal content in this enzyme were not completely pure. Furthermore, W-FDHs are extremely oxygen sensitive and often much activity is lost upon purification. Therefore the metal content of these enzymes is underestimated [44]. Based on crystallographic data of the D. gigas FDH and the Escherichia coli FDH-H, we expect that also in the S. fumaroxidans FDHs each tungsten is coordinated by two pterin cofactors and one selenocysteine [45,46]. The α2β2γ2 structure of FDH-1 then suggests that this enzyme contains 2 mol W and 2 mol Se per mol of holoenzyme, similar to the M. thermoacetica FDH [15]. Because the FDH-2 preparations used for metal analysis were pure, and much less activity of this enzyme was lost upon purification, the data presented in Table 3 suggests that this enzyme contains 1 mol W and 1 mol Se per mol of holoenzyme. The iron-sulfur centers detected in the S. fumaroxidans FDHs by EPR will be discussed below.

The presence of tungsten in enzymes has only been explored for the last two decades, with Moorella thermoacetica FDH as the first tungstoenzyme to be isolated [15]. Though there are many similarities in the properties of tungsten and molybdenum, tungsten seems to be preferred in enzymes catalyzing low potential reactions [44]. At least four of the five W-FDHs that have been isolated so far (including the two enzymes described in this study) were shown to possess CO2-reduction activity (Table 6). The D. gigas enzyme may be an exception, though to our knowledge this enzyme was never assayed for CO2-reduction [16,38]. Partially purified enzymes of Clostridium acidiurici, Clostridium formicoaceticum, as well as a second FDH in E. acidaminophilum, are also W-containing enzymes which reduce CO2[17,47–49]. The Mo-containing FDH of Wolinella succinogenes did not reduce CO2 while for most other Mo-containing FDHs isolated so far no CO2-reduction rates were reported [18,19,50–54]. Only the enzyme from Clostridium pasteurianum has been described to reduce CO2[55], but it is possible that the reduction of a substrate other than CO2 was measured considering the assay conditions used (pH 8.8 and dithiothreitol as electron donor). In summary, all formate dehydrogenases conclusively shown to perform CO2-reduction are tungsten-dependent. From these data we hypothesize that all CO2-reductases are W-containing reversible FDHs, whereas enzymes used by anaerobes to oxidize formate are Mo-containing enzymes, and probably all of these are irreversible. W-FDHs are either functional as CO2-reductases in the Wood pathway which many Clostridia explore to fix CO2, or as terminal CO2-reductases in acetogenic proton- and CO2-reducing syntrophic bacteria. We speculate that the Mo-containing enzymes evolved from W-containing CO2-reductases in organisms for which formate began to serve as an electron donor, instead of being excreted. Molybdenum is present at much higher concentrations than tungsten in most natural environments [56], and tungsten is apparently only required for FHDs that reduce CO2.

Table 6. Comparison of the Syntrophobacter fumaroxidans formate dehydrogenases to those of other organisms. Sfu, Syntrophobacter fumaroxidans (FDH-1 and FDH-2); Mth, Moorella thermoacetica; Eac, Eubacterium acidaminophilum; Dgi, Desulfovibrio gigas; Dvu, Desulfovibrio vulgaris (Hildenborough); Dde, Desulfovibrio desulfuricans. ND; not determined or reported.
SourceSubunits (kDa)StructureSpecific metalsSpecific activitya (U·mg−1)Km-formate (mm)
  1. a Formate oxidation rate, b During purification specific activities of up to 1500 U·mg−1 were measured.

Sfu189, 59, 19α2β2γ2  W+Se 7000.04
Sfu292, 33αβ  W+Se27000.01
Mth96, 76α2β22 W+2Se1100ND
Eac298, 62, 352 (αβ2γ)  W+Se 2282ND
Dgi92, 29αβ1 W  34ND
Dvu84, 27, 14αβγMo?  361.7
Dde88, 29, 16αβγ1Mo+1Se  7821

Our preliminary biochemical data suggest that in many respects the S. fumaroxidans FDH-2 is similar to the D. gigas FDH, which has been characterized more extensively [16,46]. EPR analysis of this enzyme in its fully reduced state revealed two [4Fe-4S] clusters. Preliminary crystallographic data, however, revealed besides a selenium atom coordinated to the W site, two additional [4Fe-4S] clusters. Quantification of the FDH-2 suggested that there are two [4Fe-4S] clusters in this enzyme, while at least twice as much Fe was determined by metal analysis. This suggests that perhaps this enzyme contains two additional [4Fe-4S] clusters. The absence of EPR signals originating from these additional clusters is consistent with proposed role of FDH-2 as a CO2-reductase, as the low redox potential of this reaction (E0 = −462 mV at pH 8) implies that some of the electron-relaying centers present in the enzyme should have very low redox potentials, and hence might not be reducible by dithionite. In that case, however, we would not have a suitable explanation for the fact that these additional clusters are not visible in the fully reduced spectrum of FDH-2, because Raaijmakers et al. [46] argued that they missed the additional spectrum due to loss of metal centers. The radical found in the EPR spectrum of FDH-2 most likely originates from an amino acid residue in the protein or from the pterin cofactor. Luykx et al. found that the pterin cofactor in three different molybdenum-containing aldehyde dehydrogenases produced radicals which gave rise to an isotropic S = 1/2 signal at g = 2.004 exhibiting six partially resolved hyperfine lines under nonsaturating conditions [57]. However we were unable to resolve any hyperfine interactions in the isotropic signal found for FDH-2. Therefore, we are unable to attribute the radical to a specific amino acid residue or to the pterin cofactor. Furthermore, there is a possibility that the formation of the radical in FDH-2 is an artefact due to the sample preparation or protein purification. A more extensive investigation into the formation of the radical and the EPR properties of this species is necessary to establish the nature of the radical. Our data suggest that FDH-1 is more similar to the M. thermoacetica FDH since both [2Fe-2S] and [4Fe-4S] clusters were also detected in this enzyme and the total amount of FeS centers was also unusually high.

References

  1. Top of page
  2. Abstract
  3. Experimental procedures
  4. Organism and growth conditions
  5. Enzyme purification
  6. Quantification of the formate dehydrogenase levels
  7. Enzyme assays and kinetic analyses
  8. Molecular composition of the purified formate dehydrogenases
  9. Staining of formate dehydrogenase in polyacrylamide gels
  10. N-terminal amino acid sequencing
  11. EPR spectroscopy
  12. Results
  13. Purification of the Syntrophobacter fumaroxidans formate dehydrogenases
  14. Quantification of the formate dehydrogenase levels
  15. Characterization of S. fumaroxidans formate dehydrogenases
  16. EPR spectroscopy
  17. Discussion
  18. Acknowledgements
  19. References
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