Bioconversion of CO to formate by artificially designed carbon monoxide:formate oxidoreductase in hyperthermophilic archaea

Ferredoxin-dependent metabolic engineering of electron transfer circuits has been developed to enhance redox efficiency in the field of synthetic biology, e.g., for hydrogen production and for reduction of flavoproteins or NAD(P)+. Here, we present the bioconversion of carbon monoxide (CO) gas to formate via a synthetic CO:formate oxidoreductase (CFOR), designed as an enzyme complex for direct electron transfer between non-interacting CO dehydrogenase and formate dehydrogenase using an electron-transferring Fe-S fusion protein. The CFOR-introduced Thermococcus onnurineus mutant strains showed CO-dependent formate production in vivo and in vitro. The maximum formate production rate from purified CFOR complex and specific formate productivity from the bioreactor were 2.2 ± 0.2 μmol/mg/min and 73.1 ± 29.0 mmol/g-cells/h, respectively. The CO-dependent CO2 reduction/formate production activity of synthetic CFOR was confirmed, indicating that direct electron transfer between two unrelated dehydrogenases was feasible via mediation of the FeS-FeS fusion protein.

lectron transfer is central to various essential metabolic pathways. For example, the electron transport chain and Fe-S proteins are essential constituents of the respiratory complex in all life forms on Earth. Fe-S proteins, which are involved in enzyme catalysis, regulation, maintenance of protein structure, and biological electron transfer 1,2 , include [2Fe-2S], [3Fe-4S], and [4Fe-4S] types 1, [3][4][5] . Quantum-mechanical electron transfer between Fe-S clusters could explain the maximum distance of 14 Å for physiologically relevant electron tunneling 6 . This distance has been unable to provide artificially until now and has been observed only in Fe-S proteins interacting each other in nature. Unlike other electron carriers, Fe-S proteins are used as a specific electron path, such as electric wire, so they are attractive candidates for direct electron transfer without electron loss. Ferredoxins are small (~11 kDa) soluble electron carriers that bind to proteins that contain electrons held by reduced Fe-S clusters, providing unique opportunities for engineering and synthetic biology applications [7][8][9][10][11][12][13] . Artificial fusion of Fe-S proteins, such as ferredoxin and thioredoxin-like proteins, has been performed to improve electron transfer between redox partners 7-10, [14][15][16][17] . Despite the reliable efficiency of protein fusion for facilitating electron transfer, target enzymes of fusion constructs are restricted to specific redox pairs that naturally interact with each other, such as ferredoxin and ferredoxin-dependent hydrogenase. Thus, a synthetic electron transferring path between two non-interacting redox enzymes has not yet been reported.
In this study, we attempted direct electron transfer between two different oxidoreductases, carbon monoxide dehydrogenase (CODH) and formate dehydrogenase (FDH), as a model system. CODH and FDH catalyze oxidoreduction of CO/CO 2 and formate/ CO 2 , respectively [18][19][20][21][22][23][24] . Theoretically, the overall reaction of the oxidation of CO to CO 2 with the reduction of CO 2 to formate is thermodynamically exergonic (ΔG´o = −16.5 kJ/mol). CO oxidation coupled with CO 2 reduction to formate by connecting CODH and FDH is an ideal system for monitoring electron flow between the two redox enzymes. The electric current can be easily read-out as formate, and the overall reaction does not require an additional substrate other than only CO. We chose the carboxydotrophic and formatotrophic euryarchaeota Thermococcus onnurineus NA1 as a model organism, which grows at 63°C−90°C (optimum 80°C) 25 . T. onnurineus NA1 harbors genes encoding both CODH and FDH (codhB and fdh3A, respectively) and shows high cell resistance against both CO and formate [26][27][28] . Notably, the hydrogendependent CO 2 reductase (HDCR) enzyme complex catalyzes CO 2 reduction to formate in Acetobacterium woodii using hydrogen directly or coupled CODH-ferredoxin indirectly as an electron donor 29 . However, no natural enzymes have been shown to catalyze direct CO oxidation coupled with formate production.
Here, we constructed synthetic carbon monoxide:formate oxidoreductase (CFOR) for direct electron transfer between CODH and FDH using an electron-transferring Fe-S fusion protein in T. onnurineus. The synthetic CFOR complex was purified and assayed to assess electron transfer ability in vitro as well as in vivo.

Results
Construction of CO/formate bioconversion mutants via molecular fusion of two Fe-S proteins. The redox proteins CODH and FDH from T. onnrineus NA1 were systematically engineered through molecular fusion of electron-transferring Fe-S proteins to construct a single redox complex. The codh and fdh3 gene clusters included the codhABCD and focA-fdh3ABC operons, respectively 26,30 (Supplementary Fig. 1a and Supplementary  Table 3). The codhABCD operon is responsible for CO-dependent ATP synthesis 31  Fusion of Fe-S proteins led to the formation of protein complexes associated with CODH and FDH, termed synthetic CFOR. Therefore, the fdh3B or fdh3C genes were fused directly to the codhA gene using Gibson Assembly in two possible arrangements, fdh3BC:codhA and fdh3B:codhA ( Supplementary  Fig. 1b, c). Structurally, the N-and C-termini of FdnH are located on the distal-end [4Fe-4S] cluster 22 ; thus, the distal-end [4Fe-4S] cluster at each Fe-S protein was expected to be aligned face-toface in every possible fusion combinations. Predicted models of the synthetic CFORs are presented in Supplementary Fig. 3. Next, the two flexible linkers (GGGGS) 1 and (GGGGS) 2 were designed with the fdh3BC:codhA fusion arrangement. However, (GGGGS) 3 insertion was obtained during the homologous recombination of the (GGGGS) 2 , resulting in three different lengths of linkers, which was confirmed by sequencing analysis. The fdh3BC:codhA fusion constructs pFd3CoL1C1118, pFd3CoL2C1119, and pFd3CoL3C1120 carried three different lengths of flexible linkers, i.e., (GGGGS) 1 , (GGGGS) 2 , and (GGGGS) 3 , respectively (Supplementary Table 1). Notably, the shortest linker, (GGGGS) 1 , showed the highest formate productivity (Fig. 1b, c) and was selected for fdh3B:codhA fusion.
A fosmid vector was used to facilitate cloning for chromosomal insertion of the synthetic CFOR. Insertion of the expression construct was targeted to a region of the chromosome between convergent genes TON_1126 and TON_1127, as previously described 36 . A 9-kbp DNA fragment containing the fdh3 and codh region and the P 0157 promotor-HMG cassette was inserted into the chromosome of T. onnurineus D02 by transformation of pFd3CoL1C1118, pFd3CoL2C1119, and pFd3CoL3C1120, generating the mutant strains BCF01, BCF02, and BCF03, respectively ( Supplementary Fig. 1b). Strain BCF13 was then constructed by transformation of the pFd3NStrepCoL1C1149 fosmid, which contains the fdh3B:codhA fusion with (GGGGS) 1 , into T. onnurineus D04 strain with additional deletion of the fdh3 whole gene cluster ( Supplementary Figs. 1c and 4a). An affinitypurification Strep-tag was also inserted within the operon at the N-terminus of fdh3A to allow easy purification of the synthetic CFOR enzyme at the strain BCF13 ( Supplementary Fig. 1c). Strain D05 was also constructed as a negative control for BCF13, which has no fusion linker between fdh3B and codhA (Supplementary Fig. 4b). To purify the CodhAB sub-complex, strep-tag was fused to N-terminus of CodhB, CODH catalytic subunit, and then transformed into T. onnurineus D06 strain to construct T. onnurineus D07 (Supplementary Table 1).
In summary, strains BCF01, BCF02, and BCF03 harbor Fdh3BC-CodhA fusions of different lengths of (GGGGS) 1 , (GGGGS) 2 , and (GGGGS) 3 , respectively. Strain BCF13 harbors the CFOR complex, composed of Fdh3B-CodhA fusion of (GGGGS) 1 linker. The genotype of strain D05, which was constructed to confirm the linker effect, is identical to that of strain BCF13, except that there is no linker between Fdh3B and CodhA. This strain was also used for the purification of the Fdh3AB sub-complex. Strain D07 was used for the purification of the CodhAB sub-complex.
Determination of CO-dependent cell growth and formate production. Growth and metabolic profiles of the mutant strains were then compared. Initially, cell growth, formate production, and pH change were determined from T. onnurineus BCF01, which contained the fusion of fdh3C:codhA with the shortest length of (GGGGS) 1 linker. The growth rate of this strain was similar to that of the parental strain, whereas maximum cell growth was slightly reduced (Supplementary Fig. 5a). Cumulative formate production was detected during cell growth with a final concentration of 3.2 ± 0.2 mmol/L after 60 h of incubation for the BCF01 strain, whereas formate levels were below the detection limit for the parental strain ( Supplementary Fig. 5b). Based on the overall reaction, equivalent amounts of formate and H + are produced, thereby decreasing the pH during formate production. Indeed, the pH of the cell supernatant was significantly decreased from pH 6.9 to 5.0 only in the BCF01 strain ( Supplementary  Fig. 5c). This may affect sustainable formate production and cell growth, which are optimal at pH 6.5 under CO supplementation conditions 28 . Thus, 0.1 M bis-Tris propane (pH 6.5) buffer was added to the cell growth medium to prevent the abrupt pH change due to bioconversion of CO to formate, thus stabilizing the pH without growth inhibition ( Supplementary Fig. 6).
Next, we compared the formate production ability of the remaining mutants. Cell growth was identical, and formate production was detected (Fig. 1). BCF01 showed the highest formate production (7.5 mmol/L) after 60 h of incubation (Fig. 1b). The relative formate productivities of BCF02 and BCF03 at the final time point were determined to be 78% and 84%, respectively, compared with BCF01 (Fig. 1c). The (GGGGS) 1 linker showed the highest formate productivity; however, the effect on the electron transfer efficiency was insignificant. BCF13, carrying the fdh3B:codhA fusion with the (GGGGS) 1 linker, showed 6.6 ± 2.1 mmol/L formate production under CO-supplemented cell growth within 24 h (Fig. 1d), which was higher than that of BCF01 (4.3 ± 0.4 mmol/L formate) at the same time. Cell growth was similar to the other strains. Therefore, all subsequent experiments were conducted using the BCF13 strain. Formate production from the cell suspension was also investigated in serum vials using the strain D05 (without fusion linker) and BCF13 (with fusion linker) at an OD 600 of about 0.5, incubated at 80°C in the presence of CO gas with 2 bar (gauge pressure) CO/CO 2 (50:50 v/v) or CO/N 2 (50:50 v/v) mix gas. Activity and stability of the cell suspensions were confirmed by H 2 productivity that showed similar values among strains and gas conditions (Fig. 1f). In the BCF13, 3.0 ± 0.27 mmol/L formate was produced after 60 min incubation under the CO/CO 2 mix gas (Fig. 1e). In contrast, when the headspace was filled with CO/N 2 , only 0.3 ± 0.04 mmol/L formate was produced due to the low CO 2 partial pressure ( Fig. 1e and Supplementary Fig. 7). Although the CO oxidation reaction provides the equivalent CO 2 requirement for the formate production, the additionally supplemented CO 2 enhances the CO 2 reduction/formate production reaction. In the D05, formate production was detected under CO/CO 2 mix gas as a concentration of 0.1 ± 0.007 mmol/L at 60 min, which is 30-fold lower than the BCF13 (Fig. 1e). The results indicate that electron transfer between CODH and FDH modules is achievable just by overexpression of the enzymes but extremely enhanced by a flexible fusion of FeS-FeS in the synthetic CFOR complex.
Purification of the synthetic CFOR complex. We then purified the CFOR enzyme complex isolated from strain BCF13 grown in a fed-batch bioreactor with CO-supplemented MM1 medium to activate the expression of genes controlled by the strong promoter P 0157 , which induces robust transcription and translation under CO-supplemented growth conditions 37 . The CFOR complex was purified using a strep-tag fused to the N-terminus of the FDH catalytic subunit, Fdh3A, and then analyzed by SDS-PAGE. Fdh3A, CodhB, and Fdh3B-CodhA fusion subunits, were present, with apparent molecular masses of 76, 67, and 42 kDa, indicating that the Fdh3B-CodhA fusion protein spontaneously bound to CodhB and Fdh3A to form the CFOR complex ( Fig. 2a and Supplementary Fig. 8). Protein bands consistent with the calculated molecular weights from deduced amino acid sequences were observed for all three subunits (Supplementary Table 3). The calculated size of Fdh3B-CodhA was 43,722 Da. The CFOR complex was further purified using size-exclusion chromatography, and the CFOR complex from gel filtration was eluted as a single major peak with an apparent mass of around 488 kDa (Fig. 2b, c). The three major bands of purified protein were identified by LC-MS/MS analysis by bands cut from the SDS-PAGE gel ( Fig. 2a  CodhB, respectively. SDS-PAGE showed Fe-S small subunit, Fdh3B and CodhA, with an apparent molecular mass of 18 kDa and 24 kDa, respectively (Fig. 2a). Based on the molecular weight of CFOR and the information from the protein structure of CODH and FDH 22,41 , it seems reasonable to conclude that CFOR has an octameric structure of 4CodhB, 2Fdh3A, and 2Fdh3B-CodhA fusion protein, equivalent to a dimer of 2CodhB, 1Fdh3A, and 1Fdh3B-CodhA tetramers. Therefore the molar ratio of the sub-complexes, Fdh3AB and CodhAB in CFOR, was calculated as 2.2 and 3.8, respectively, which is similar to the protein band intensity of those of CFOR subunits (Supplementary Tables 5, 6).
Catalytic properties of the CFOR enzyme complex. The activities of CODH and/or FDH from the purified CFOR, CodhAB, and Fdh3AB enzymes were determined individually using the methyl viologen assay method. The specific CO oxidation activity was determined to be the same level in isolated CFOR and Cod-hAB as 1,548.5 ± 366.9 μmol/mg/min and 1,585.6 ± 21.8 μmol/mg/ min, respectively. In contrast, there was a huge difference in FDH activity between CFOR and Fdh3AB. The specific formate oxidation activity was determined as 17.1 ± 0.3 μmol/mg/min and 2,139.7 ± 54.4 μmol/mg/min from the CFOR and Fdh3AB, respectively (Table 1). Why the FDH activity of CFOR complex turns to be extremely low is unclear, but complex formation by protein fusion is thought to be one of the causes of it. In the CFOR, the specific activity of FDH was lower than that of CODH; therefore, the reaction of FDH was expected to be a rate limiting factor for the formate production in the overall CFOR reaction. To confirm direct electron transfer by the Fe-S fusion protein, we investigated whether the purified CFOR could catalyze CO oxidation/formate production without additional electron carriers. The enzyme indeed catalyzed formate production from CO/CO 2 (50:50 v/v) mix gas with a maximum specific activity of 2.2 ± 0.2 μmol/mg/min at 20 min ( Fig. 3 and Table 1). However, formate was below the detection limit under conditions with 100% CO or without CFOR enzyme, which result corresponds to that of the cell suspension experiment (Fig. 1d). In the reaction, CO 2 requirement is equilibrium with CO 2 generation by CO oxidation; therefore, it could be considered that CO 2 partial pressure is not related to the formate production reaction. However, CO 2 partial pressure is closely related to the initial formate production rate. When the reaction begins with only CO, generated CO 2 by CO oxidation at the CODH active site will be diffused in the solution immediately, rather than reduced at the FDH active site, because a specific CO 2 channel is absent in the CFOR system. Therefore, an appropriate CO 2 partial pressure is needed for the rapid CO 2 reduction at the FDH. As a control experiment for the formate production by CFOR, purified sub-complexes Fdh3AB (18.3 ug) and CodhAB (31.7 ug) were mixed corresponding to 50 ug of CFOR according to the calculated molar ratio (Supplementary Table 5) and then demonstrated the formate production assay under the same condition. After 60 min of reaction, only 0.051 ± 0.05 mM of formate was detected in the Fdh3AB and CodhAB sub-complex mixed sample, whereas 2.47 ± 0.51 mM, 49fold higher, was detected in the CFOR complex (Fig. 3). A Fdh3-Codh mixture containing 198 pmol of methyl viologen as the electron carrier corresponding to the FdhB-CodhA fusion protein was also tested, and 0.017 ± 0.015 mM of formate production was detected in 60 min. The results demonstrated that the simple, flexible fusion of two Fe-S proteins enabled electron transfer between them, leading to the formation of functional enzyme by an assembly of non-interacting redox enzymes.
Bioconversion of CO to formate in a bioreactor. The formate production potential of the strain was tested in a bioreactor where ARTICLE COMMUNICATIONS BIOLOGY | https://doi.org/10.1038/s42003-022-03513-7 100% CO was continuously fed with a flowrate of 0.02-0.122 vvm (CO volume/working volume/min). Table 2 summarizes thebioreactor parameters for the T. onnurineus BCF13 strain. Formate production was detected at a concentration of 56.4 ± 6.4 mmol/L after fermentation for 6 h (Fig. 4b). The formate production rate and maximum specific formate productivity were calculated as 13.1 ± 0.9 mmol/L/h and 73.1 ± 29.0 mmol/gcells/h, respectively. Recently CO-dependent formate production by coupling of CODH, ferredoxin, and HDCR was reported using A. woodii and Thermoanaerobacter kivui as a whole-cell biocatalyst; the formate production rate was 0.28 mmol/L/h 42 . The specific rates were determined as 1.44 and 1.34 mmol/g/h for A. woodii Δrnf and T. kivui, respectively 43 . T. onnurineus NA1 and its derivatives mutants used in this study are a basic hydrogenogenic carboxydotroph that can grow on CO as an energy source via the CO-dependent respiratory gene cluster codh-mch-mnh3 26,28,30 . Thus, the BCF13 strain showed carboxydotrophic properties, such as H 2 and CO 2 production (Figs. 1f and 4c), and could grow under 100% CO conditions with the maximum specific growth rate (μ max ) of 0.621 ± 0.051 h −1 (Fig. 4a). This spontaneous CO 2 production by the CO-dependent respiration enhances the formate production in the bioreactor. Therefore, T. onnurineus BCF13 could be used as an industrial microorganism for the production of H 2 and formate simultaneously from CO.

Discussion
Fe-S proteins involved in the electron transfer chain have been well characterized, both structurally and functionally. Fe-S proteins in many oxidoreductase complexes are typically associated   with a large subunit that has catalytic redox activity and forms an enzyme complex, such as respiratory complex I, FDH-N and formate hydrogenlyase in E. coli 22,32,44 . However, direct electron transfer between CODH and FDH has not yet been found in nature. Thus, if one could construct an electron transfer system between these two oxidoreductases, it could serve as a universal electron transfer system. Accordingly, we generated two different fusion combinations, Fdh3BC-CodhA and Fdh3B-CodhA to, create an electron transfer path, and as a result, CO 2 reduction/ formate production reaction by electron transport was observed in all combinations. The small subunits CodhA and Fdh3B specifically interacted with their catalytic large subunits. Therefore, the molecular fusion of Fe-S proteins may spontaneously mediate the formation of a CODH-FDH protein complex (Fig. 2). A significant amount of formate production was only detected from the synthetic CFOR enzyme complex, but not for the individual mixtures of sub-complexes supplemented with or without methyl viologen as an electron carrier (Fig. 3). The results suggest that the electrons from CO oxidation by the CODH are not shared with other electron acceptors and are transferred directly to the FDH, which leads to a concentrated reducing power showing the highest CO 2 reduction and formate production rate reported so far. According to the electron tunneling theory, the maximum distance between the distal end [Fe-S] clusters at each protein must provide a distance of at least 14 Å for electron transfer, indicating that tight-binding and a sophisticated rearrangement of the two proteins are essential, and which was achieved by the flexible linker in the CFOR. Some potential possibilities can be associated with this phenomenon. First, the two sub-complexes have been connected by a flexible linker peptides which allows for motility of the connecting proteins and can move randomly during the reaction 45 . Electrons may transfer if the distal end [Fe-S] clusters of each Fe-S protein coincidently collide by the random motion at an uncertain frequency. Second, the incorporation of Ser residue in the flexible linker can maintain the stability of the linker in aqueous solutions by forming hydrogen bonds with the water molecules 46 . Electron transfer may be induced by these hydrogen bonds mediating rearrangement between the distal [Fe-S] clusters. However, the principles of direct electron transfer by Fe-S fusion protein are unclear, and the validations remain further.
The FDH activity of the CFOR complex significantly decreased than the native Fdh3AB sub-complex (Table 1). It is not clear why the FDH activity is different between the two complexes, but it may result from the conformational change of Fdh3A by protein fusion. Some FDHs from bacteria, including Desulfovibrio alaskensis, form heterodimeric (αβ) 2 structures 47 . If Fdh3AB also formed such (αβ) 2 structure, the Fdh3B-CodhAB fusion protein may cause a conformational change in the catalytic moiety, resulting in a lack of FDH activity. Indeed, the CFOR was predicted as a heterodimeric (Fdh3AB-CodhAB 2 ) 2 , which can be due to the dimerization of Fdh3AB. To enhance FDH activity, various construction of CFOR using monomeric FDHs and evaluations of enzymatic activity could be necessary.
This work focused on the direct electron transfer between two unrelated redox enzymes by a fusion of Fe-S proteins as an electron path. Using this electron path, we connected noninteracting carbon monoxide dehydrogenase and formate dehydrogenase, constructing a synthetic CFOR as a single functional enzyme complex. The mutant strain BCF13, harbored CFOR encoding genes, showed efficient CO conversion/formate production ability of 73.1 ± 29.0 mmol/g-cells/h. The purified CFOR was also exhibited 2.2 ± 0.2 μmol/mg/min of specific formate productivity from CO. Overall, our results provide some insight into the synthesis of an electron path using the simple fusion of Fe-S proteins, which can be applied to various combinations of redox enzymes for efficient CO 2 reduction and production of value-added chemicals.

Methods
Strains and cell culture conditions. T. onnurineus NA1 wild-type and mutant strains were routinely cultured in modified medium 1 (MM1) 31,48 containing 4 g/L of yeast extract (BD Bioscience, San Jose, USA) and 4x Holden's trace element/Fe-EDTA solution at 80°C. All procedures for the cultivation of T. onnurineus strains were carried out in an anaerobic chamber (Coy Laboratory Products, Grass Lake, USA). For the cultures in serum bottles, bis-Tris propane (pH 6.5) buffer was additionally supplemented in the medium to a final concentration of 0.1 M, and the  headspaces were filled with 100% CO (MMC) was provided to support the growth of wild-type and mutant strains. The serum bottles were sealed with bromobutyl rubber stoppers and aluminum crimp caps. All procedures were carried out under strictly anaerobic conditions. For the pH-stat batch culture, T. onnurineus strain BCF13 was serially cultured in a 150 ml serum bottle and 7 L bioreactors (Fermentec, Cheongwon, South Korea); the working volumes of which were 80 ml and 5 L of MM1 medium, respectively, at 80°C. The bioreactors were sparged with pure argon gas (99.999%) through a microsparger. The agitation speed was 500 rpm, and the pH was controlled at 6.2 using 2 M KOH in 3.5% NaCl. The inlet gas of 100% CO was supplied by using a mass flow controller (MKPrecision, Seoul, South Korea) at feeding rates of 100−610 ml/min. E. coli EPI300 TM -T1 R (Epicentre Biotechnologies, Madison, USA) strain was used for fosmid based molecular cloning purposes. Fosmid containing E. coli clones were cultured in LB medium containing 12.5 ug/ml chloramphenicol. General molecular biology manipulations and microbiological experiments were carried out by standard methods 49 .
Cloning and construction of the CFOR expression vector. The cloning strains, plasmids and fosmids used in this study are listed in Supplementary Table 1. The fdh3 gene cluster deletion mutant (strain D04) was constructed by the previously used gene disruption system in T. onnurineus NA1 36 (Supplementary Fig. 4a). To construct fosmid vector backbone, previously constructed complementary insertion site (Left_arm (TON1128-TON_1127) region-P 0157 promotor-HMG cassette-Right_arm (TON_1126) region) 36 was amplified by PCR using the primers listed in Supplementary Table 2. The amplicon and fosmid vector pCC1FOS (Epicentre Biotechnologies, Madison, USA) were assembled into a single vector, pNA1-comFosC1096 (Supplementary Table 1), using a SLIC method 50 . PCR products of the T. onnurineus NA1 Fdh3 encoding gene region (focA-fdh3ABC or focA-fdh3AB), Codh encoding gene region (codhABCD), and fosmid vector backbone (pNA1comFosC1096 with AvrII enzyme digestion) were assembled into a single vector using Gibson Assembly Master Mix (New England Biolabs, Ipswich, USA). The fusion targeting Fe-S protein-encoding genes, fdh3B or fdh3C, and codhA from T. onnurineus NA1 were fused using a homologous recombination event with 26 bp complementary PCR primers to generate (GGGGS) 1-2 flexible linker sequence during the gene assembly reaction by the Gibson Assembly method. The 3'-end of fdh3B or fdh3C gene lacking its stop codon was fused to the 5'-end of codhA gene lacking its start codon mediated by (GGGGS) 1-2 linker (Supplementary Fig. 1). The sequences of the fusion genes were verified by DNA sequencing. Transformations of T. onnurineus strains with the constructed fosmid and the confirmation of transformants were performed by PCR 28 . The primers used to create the fusion genes are listed in Supplementary Table 2.
Analytical methods. Cell growth was monitored by measuring optical density at 600 nm (OD 600 ) with a spectrophotometer (Eppendorf, Hamburg, Germany). The unit value of OD 600 corresponded to 0.361 g/L (dry cell weight) as previously determined 37 . The amounts of CO, CO 2 , and H 2 gas were measured using a gas chromatograph (GC; YL Instrument Co., Anyang, South Korea) equipped with a Molsieve 5A column (Supelco, Bellefonte, PA), a Porapak N column (Supelco, Pennsylvania, USA), a thermal conductivity detector, and a flame ionization detector 37 . Formate was measured by high-performance liquid chromatography (HPLC; YL Instrument Co., Anyang, South Korea) with a Shodex RSpak KC-811 column (Showa Denko, Kanagawa, Japan). Ultrapure water containing 0.1% (v/v) phosphoric acid was used as the mobile phase at a flow rate of 1.0 ml/min. All samples were prepared with 1 ml of culture broth and centrifuged to remove cell debris at 4°C, 13,480 × g for 5 min. The supernatants were purified with a syringe filter and analyzed by HPLC.
Cell suspension experiment. To prepare cell suspensions, T. onnurineus strain BCF13 was anaerobically cultured in a 7 L fermentor with a working volume of 3 L as described above. At the end of the culture, the cells were harvested by centrifugation at 5523 × g for 30 min at 20°C and then washed two times with an anaerobic modified PBS (600 mM NaCl, 2.7 mM KCl, 10 mM Na 2 HPO 4 , 2 mM KH 2 PO 4 , and 2 mM DTT). Finally, cells were resuspended in the same buffer at cell densities of OD 600 = 0.5. Four milliliters of cell suspensions were transferred to a 20 ml serum vial under a headspace of CO/CO 2 (50:50 v/v) mix gas at about 2 bar (gauge pressure), or CO/N 2 (50:50 v/v) mix gas at about 2 bar (gauge pressure), respectively. The cell suspensions were incubated at 80°C, and then the formate concentration and headspace gas composition were determined by HPLC and GC, respectively.
Purification of enzymes. To purify CFOR enzyme complex, typically 2 to 4 g (wet weight) of fed-batch cultured T. onnurineus strain BCF13 cell pellets were harvested and washed with the modified PBS and then resuspended in buffer W (100 mM Tris-HCl, 150 mM NaCl, pH 8.0). Cells were then disrupted by sonication on ice, and cell debris was removed by centrifugation (15,000 × g for 40 min at 4°C). Affinity column purification was carried out following the manufacturer's protocols with a Strep-Tactin system (IBA-Lifsciences, Göttingen, Germany). The molecular weight and additional purification of CFOR complex were determined by analyzing the purified protein on a calibrated Superdex 200 10/300 GL column equilibrated buffer W using fast protein liquid chromatography (Äkta FPLC System, Amersham Biosciences). The column was calibrated by using these standards: thiroglobulin (669 kDa), apoferritin (443 kDa), beta-amylase (200 kDa), albumin (66 kDa) and carbonic anhydrase (29 kDa). Strep-tag purified protein was loaded and eluted at a flow rate of 0.5 ml/min, then the fractions were selectively collected at about 488 kDa of a single peak. The above procedures were carried out under anaerobic conditions. The purified proteins by size exclusion chromatography were used for enzyme assay, and examined via sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) according to the standard methods. The major bands were identified by LC-MS/MS analysis service (Yonsei Proteome Research Center, Seoul, Korea).
Enzyme assays. CODH activity was assayed at 80°C by a colorimetric assay with methyl viologen (MV) as an electron acceptor (ε 578 = 9.7 mM/cm at 578 nm) 51 , and CO as an electron donor. The assay was conducted with 4.4 ng of purified CFOR complex in 2 ml volume of sodium phosphate buffer (50 mM sodium phosphate, and 2 mM DTT, pH 7.5) containing 5 mM MV in a 4 ml serumstoppered glass cuvette. 100% CO gas was purged in the headspace at about 2 bar (gauge pressure), and then the reaction was initiated by incubation of the reaction mixture at 80°C. Formate dehydrogenase activity was assayed using 360 ng of purified CFOR complex under the same conditions except that CO was replaced by 10 mM sodium formate. The formate production assay was carried out in 2 ml volume of 50 mM sodium phosphate buffer (pH 7.5) containing 2 mM DTT and 50 ug of purified CFOR in a 20 ml serum-stoppered vial. CO gas was purged in the headspace at about 2 bar (gauge pressure) of CO/CO 2 (50:50 v/v) mix gas, or at about 0.5 bar of 100% CO gas. The reaction was initiated by incubation at 80°C. Formate in reaction mixture was determined by HPLC.
Thermodynamic calculation. The biological standard Gibbs energy value (ΔG´o) was calculated by the Nernst equation using values of the standard Gibbs energy (ΔG o ) reported by Amend and Shock 52 .
Statistics and reproducibility. In all figures, error bars represent the standard deviation of the mean value. To determine product formation from the mutant strains and the CFOR enzyme using HPLC or GC, the number of replicates performed at least three biological replicates (n = 3) in every experiment (shown as mean ± s.d.), were measured to reveal a similar level of the product.