Significance High carbon yield in the bioengineering of heterotrophic bacteria is hindered by carbon loss to CO 2 production. We provide evidence showing Clostridium thermocellum, a cellulose-degrading bacterium and a model consolidated bioprocessing organism, can fix CO 2 while growing predominantly on cellobiose, a cellulose-derived disaccharide. By adding 13C-bicarbonate to the bacterial culture and tracking 13C-labeled metabolites, we discovered active reductive one-carbon (C1) metabolism in this bacterium. We further identified critical enzymes responsible for fixing CO 2 and channeling the fixed carbon to the C1 metabolic pathway. Our findings pave the way to future engineering of this bacterium to use cellulose and CO 2 simultaneously as a means to improve microbial carbon efficiency that is constrained by theoretical limitation and to reduce CO 2 in the environment.

Abstract Clostridium thermocellum can ferment cellulosic biomass to formate and other end products, including CO 2 . This organism lacks formate dehydrogenase (Fdh), which catalyzes the reduction of CO 2 to formate. However, feeding the bacterium 13C-bicarbonate and cellobiose followed by NMR analysis showed the production of 13C-formate in C. thermocellum culture, indicating the presence of an uncharacterized pathway capable of converting CO 2 to formate. Combining genomic and experimental data, we demonstrated that the conversion of CO 2 to formate serves as a CO 2 entry point into the reductive one-carbon (C1) metabolism, and internalizes CO 2 via two biochemical reactions: the reversed pyruvate:ferredoxin oxidoreductase (rPFOR), which incorporates CO 2 using acetyl-CoA as a substrate and generates pyruvate, and pyruvate-formate lyase (PFL) converting pyruvate to formate and acetyl-CoA. We analyzed the labeling patterns of proteinogenic amino acids in individual deletions of all five putative PFOR mutants and in a PFL deletion mutant. We identified two enzymes acting as rPFOR, confirmed the dual activities of rPFOR and PFL crucial for CO 2 uptake, and provided physical evidence of a distinct in vivo “rPFOR-PFL shunt” to reduce CO 2 to formate while circumventing the lack of Fdh. Such a pathway precedes CO 2 fixation via the reductive C1 metabolic pathway in C. thermocellum. These findings demonstrated the metabolic versatility of C. thermocellum, which is thought of as primarily a cellulosic heterotroph but is shown here to be endowed with the ability to fix CO 2 as well.

The gram-positive Clostridium thermocellum is a thermophilic and strict anaerobic bacterium. It has gained a great amount of interest due to its cellulolytic abilities. By taking advantage of an extracellular cellulase system called the cellulosome (1), C. thermocellum can depolymerize cellulose into soluble oligosaccharides. The latter are further transported into the cells and fermented through its glycolytic pathway to pyruvate, the precursor to an array of fermentation products (e.g., H 2 , formate, lactate, acetate, ethanol, secreted amino acids) (2, 3). This capability makes C. thermocellum an attractive candidate for consolidated bioprocessing of lignocellulosic biomass, a process configuration that directly converts plant biomass into biofuels and chemicals without separate additions of enzymes (4). To date, there have been numerous works describing the molecular and genetic details of the cellulolytic system and fermentation pathways in C. thermocellum (5⇓⇓⇓–9), whereas other metabolic characteristics of this species have not been adequately understood.

Currently, few details regarding inorganic carbon utilization are known for C. thermocellum. However, several investigators routinely add bicarbonate, a dissolved form of CO 2 , into the medium to promote C. thermocellum growth (6, 10⇓–12), implying its ability to use CO 2 . This observation raises the questions of how CO 2 plays a role in promoting C. thermocellum growth (13), and whether CO 2 is incorporated into the metabolism. There is limited information on pathways relating to CO 2 uptake thus far. One proposed pathway involving CO 2 utilization in C. thermocellum is the malate shunt and its variant pathway (7, 14), in which CO 2 is first incorporated by phosphoenolpyruvate carboxykinase (PEPCK) to form oxaloacetate and then is released either by malic enzyme or via oxaloacetate decarboxylase complex, yielding pyruvate. However, other pathways capable of carbon fixation may also exist but remain uncharacterized.

In recent years, isotopic tracer experiments followed by mass spectrometry (MS) (15, 16) or NMR measurements (17) have emerged as powerful tools for metabolic analysis. This methodology traces the transit of isotopic atoms throughout the biochemical network and sheds light on active reactions as well as molecular fluxes in vivo. Coupled with classic genetic manipulations, this technology has been successfully applied to provide insights into novel metabolic pathways in bacteria (18), yet it has not been used to study C. thermocellum.

In several species of Clostridium, the one-carbon (C1) metabolism is initiated from CO 2 reduction to become formate via formate dehydrogenase (Fdh) (19). This pathway provides C1 units for many crucial reactions, including amino acids and purine biosynthesis. Formate serves as a precursor to the folate biosynthesis pathway that forms the “methyl” branch of the Wood–Ljungdahl pathway, yielding acetyl-CoA from CO 2 (19). However, C. thermocellum lacks the first key enzyme, Fdh, based on genome information. In the present study, we investigate CO 2 -fixing pathway(s) in C. thermocellum involving C1 metabolism by using 13C-tracer experiments. We measure the isotopomers of fermentation products and proteinogenic amino acids using NMR and GC/MS, and uncover a route C. thermocellum employs to fix CO 2 when grown in a primarily heterotrophic mode supplemented with sodium bicarbonate.

SI Results rPFOR Activity, and Not Label Exchange, Contributed to Pyruvate Labeling. To confirm rPFOR activity as the main mechanism contributing to the labeled pyruvate, and not the label exchange reaction, we fed Clostridium thermocellum [2-13C]acetate (40 mM) supplemented with unlabeled bicarbonate (20 mM) when the bacteria were grown primarily in cellobiose (14.6 mM or 5 g/L). Our results showed labeled pyruvate at the expected position (carbon at position 3) from [2-13C]acetate feeding in both the control strain (∆hpt) and the PFL inactivated mutant (∆hpt∆pflA). These results also ruled out the possibility of labeled pyruvate transiting from acetyl-CoA via reversed PFL activity. The observed labeled carbon at position 3 of pyruvate is independent of the label exchange carbon specific to the carboxyl group. We further analyzed the labeling pattern of acetyl-CoA using carbons at positions 1 and 2 (C1–2) of leucine as a proxy during [2-13C]acetate feeding in ∆hpt and ∆hpt∆pflA following the same experimental setup described. Briefly, C1–2 of N-tert-butyldimethylsilyl-N-methyltrifluoroacetamide (TBDMS)–derivatized leucine can be fragmented by GC/MS and is directly derived from acetyl-CoA. In summary, results in Fig. 3C showed that the label fraction in acetyl-CoA is 31.6% in ∆hpt∆pflA and the corresponding label fraction in pyruvate is 13.4% (in ∆hpt∆pflA). When dividing 13.4% by 31.6%, we get 0.42. The number 0.42 represents the labeled fraction transited to pyruvate from acetyl-CoA via the rPFOR activity by feeding 13C-acetate, which closely matches the fraction labeled in pyruvate by feeding 13C-bicarbonate reported in the main text, which is 43% in wild type (∆hpt; Fig. 3A). Our data consistently and clearly showed the rPFOR activity as the main mechanism contributing to pyruvate labeling from CO 2 fixation, and not label exchange. Effects of PFL KO on Serine Labeling. To investigate how PFL deficiency affects carbon flux toward C1 metabolism, we compared the labeling pattern of isotopomers in both ∆hpt∆pflA and wild type by feeding 13C-formate (20 mM) and 13C-bicarbonate (20 mM), respectively (Fig. 5). In either case, unlabeled cellobiose (14.6 mM) was added to the cell culture. For 13C-formate feeding experiments, results showed that Ser1–3 and Ser2–3 fragments in both strains have a significant portion (15–26%) of the molecules labeled on C1 (Fig. 5A). To the contrary, Ser1–2 in ∆hpt∆pflA remained nearly unlabeled and only 6% of Ser1-2 molecules in wild type had one 13C-carbon, which was probably a result of the rPFL activity. We therefore deduced that the 13C-carbon propagated from 13C-formate is mainly at position 3 of Ser (hydroxylmethylene group), consistent with our constructed de novo C1 metabolic pathway toward serine biosynthesis (Fig. 2). Effects of PFL KO on Methionine Labeling. With the addition of 13C-formate, one carbon in Met is expected to be traced back to the methyl group in methyl-THF downstream of formate according to the biochemical reactions depicted in Fig. 2. As to the 13C-bicarbonate labeling experiments in ∆hpt∆pflA, we observed a lower extent of fractional labeling of methionine in the ∆hpt∆pflA mutant than in the wild type (Fig. 5B), implying impaired reductive C1 metabolism. It should be noted that more than one carbon can be labeled in methionine molecules because methionine is not solely synthesized from the C1 metabolic pathway. For instance, in addition to the methyl group from the C1 pathway, other labeled carbons may be derived from the malate shunt, in which oxaloacetate is a carboxylated intermediate serving as the precursor for C1–4 in methionine.

Discussion This study reveals that during cellobiose metabolism, C. thermocellum also assimilates CO 2 via a reductive C1 metabolic pathway that is initiated by the dual actions of reversed PFOR and PFL. PFOR has been greatly considered in the forward direction in C. thermocellum, catalyzing the oxidative decarboxylation of pyruvate to acetyl-CoA and CO 2 and generating reduced ferredoxin (Fd red ). The PFOR activity in C. thermocellum has been associated with the oxidative decarboxylation of pyruvate to acetyl-CoA. Uyeda and Rabinowitz (24, 25) and Raeburn and Rabinowitz (26) demonstrated in vitro that low potential electron donors, such as Fd red , can drive PFOR to catalyze the reductive carboxylation of acetyl-CoA as a pyruvate synthase. The rPFOR serves as a key enzyme for autotrophic CO 2 fixation via the Wood–Ljungdahl pathway in acetogens (27). In this work, the in vivo isotopomer data measured postfeeding C. thermocellum 13C-biocarbonate and [2-13C]acetate double-confirmed rPFOR activity and revealed that rPFOR activity accounted for over 40% of the labeled carbon in pyruvate. Measurements in individual PFOR mutants further identified rPFOR activity predominantly catalyzed by Clo1313_0020-0023 and Clo1313_1353-1356. Consistent with previous global expression studies, Clo1313_0020-0023 is the most highly expressed PFOR based on proteomics (28) and transcriptomics (29) analyses. Clo1313_1353-1356 is the third most highly expressed PFOR based on proteomics data, but it has a low transcript level based on microarray data, and the difference could be due to different growth conditions in these studies. By feeding 13C-bicarbonate to the PFL-inactivated mutant and the wild type, we confirmed the incorporation of these tracers into formate (Fig. 1), lactate (Fig. 3A), and the downstream amino acids linked to C1 metabolism (Figs. 3 and 5). We provided multiple lines of evidence showing that the fixed CO 2 was channeled to the C1 metabolism through PFL, which could operate reversibly (21, 30) and contribute to the biosynthesis of essential amino acids, including serine and methionine. In addition, the reductive C1 pathway mediated by PFL may play a critical role in nucleotide biosynthesis (reconstructed purine biosynthesis pathway in C. thermocellum shown in Fig. S6). The impaired growth observed in PFL-inactivated mutant is consistent with a previous finding (12) and underscores PFL’s importance. A transcriptome study reported that folate-dependent enzymes are in the most highly expressed category, which also suggests an active C1 metabolism in vivo (31). Fig. S6. Reconstruction of purine biosynthetic pathway in C. thermocellum DSM1313. Dotted lines indicate multistep catalysis. Putative genes involved in each step are present in gene abbreviations with locus numbers. Carbons from C1 metabolism are labeled in red. ADSL, adenylosuccinate lyase; AIRC, 5-aminoimidizole ribonucleotide carboxylase; AIRS, 5-aminoimidizole ribonucleotide synthetase; ATIC, phosphoribosylaminoimidazolecarboxamide formyltransferase/IMP cyclohydrolase; GARS, glycinamide ribonucleotide synthetase; GARTFase, phosphoribosylglycinamide formyltransferase; IGPS, imidazole glycerol phosphate synthase; IMP, inosinate mononucleotide phosphate; PFAS, phosphoribosylformylglycinamidine synthase; PPAT, phosphoribosyl pyrophosphate amidotransferase; PRPS1, phosphoribosyl pyrophosphate synthetase 1. Carbon loss in the form of CO 2 from the conversion of pyruvate to acetyl-CoA across a wide range of bacterial heterotrophs reduces the theoretical maximum yield of a carbon product of interest per carbon consumed (carbon efficiency) downstream of acetyl-CoA. A synthetic nonoxidative glycolysis has been shown to bypass the loss of CO 2 in the conversion of pyruvate to acetyl-CoA (32). The rPFOR activity shown here allows some of the lost CO 2 to be recaptured into pyruvate, a critical branch point and precursor to numerous natural and nonnatural products. Based on our data, CO 2 utilization has increased the titers of organic carbon compounds (Table 1) by incorporating CO 2 into the biomass formula, as detected in proteinogenic serine and methionine, and, as such, supplementing cell growth. It is interesting that C. thermocellum uses the rPFOR-PFL shunt for CO 2 uptake in the absence of Fdh. We speculate that although pyruvate is a key branching point in central carbon metabolism and a key node connecting glycolysis and the C1 metabolic pathway, the rPFOR-PFL route for CO 2 uptake may provide added metabolic flexibility and versatility to cells in response to environmental changes affecting their carbon and energy (redox) needs. To gain insights into how widespread the rPFOR-PFL shunt could be for CO 2 uptake, we surveyed the presence of PFOR, PFL, and Fdh in 67 sequenced Clostridium species. We discovered that only 20 species have Fdh, whereas 51 species have both PFOR and PFL (Fig. S7). Although the in vivo reversibility of PFOR can be organism- and/or growth condition-specific, and information as to whether these bacteria can use inorganic carbon during organic substrate feeding may not be readily available, the presence of these genes renders it genetically feasible. Fig. S7. Summary of the survey results identifying the presence of genes encoding for PFOR, PFL, and in 67 sequenced Clostridium species. It should be noted that carbon fixation via the rPFOR-PFL shunt requires Fd red as an electron donor. On the other hand, PFOR functioning in the oxidative direction serves as a major pathway for generating Fd red to yield reduced biofuels and chemicals. Thus, the metabolic flux toward biosynthesis may be a trade-off of the flux toward electron generation. We speculate that CO 2 enhances growth by regenerating oxidized Fd, which, in turn, provides a driving force for faster cellobiose consumption. Feeding CO 2 may also enhance the “transient” CO 2 fixation via the malate shunt, with GTP produced via PEPCK promoting C. thermocellum growth. The biosynthesis vs. energy requirement at the pyruvate node needs to be evaluated comprehensively with other major redox pathways (i.e., H 2 production) to enhance the carbon economy without causing redox imbalance. To satisfy our curiosity, we tested C. thermocellum autotrophic growth by feeding either bicarbonate plus 100% H 2 or an 80% H 2 /20% (vol/vol) CO 2 gas mixture, but detected no growth (Fig. S8). Nevertheless, the rPFOR-PFL shunt uncovered in this study may have important ramifications in the metabolic plasticity of a cellulose degrader and in other heterotrophs, and hence warrants more in-depth investigations. Fig. S8. Isotopomer analysis of proteinogenic alanine in Δhpt grown on 50 mM 13C-sodium bicarbonate with 100% argon (Ar) gas in the headspace (A) and 100% H 2 gas in the headspace (B). No growth was observed, and the OD 600 of the culture grown in either Ar or H 2 gas dropped from 0.46 to 0.28 (in Ar gas) and from 0.43 to 0.29 (in H 2 gas) 24 h postinoculation. H 2 gas in the headspace remained relatively unchanged. (C) Summary of OD 600 , H 2 , and CO 2 in the headspace at t 0 and 24 h after feeding wild-type (Δhpt) 80% H 2 and 20% CO 2 gas mixture in sealed tubes. This strain has been defined previously. R.A., relative abundance.

Materials and Methods A more complete discussion is provided elsewhere (15, 16, 23, 33⇓–35), as well as in SI Methods. Unless otherwise noted, C. thermocellum DSM1313-derived strains, ∆hpt (hpt encoding for hypoxanthine phosphoribosyl transferase is used for counterselection), and other mutants were routinely grown anaerobically at 55 °C on 5 g/L cellobiose. The hpt gene deleted strain is used as the wild type for the studies herein. To test the inorganic carbon utilization, 20 mM sodium bicarbonate was added to cell cultures also fed with cellobiose. We used the DSM122-defined medium referred to as CTFUD medium (36). Full details on strain construction, fermentation conditions, and 13C-tracer experiments are provided in SI Methods.

SI Methods Strains, Culture Conditions, and Medium. Unless otherwise noted, C. thermocellum DSM1313-derived strains and mutants were routinely grown anaerobically at 55 °C on 5 g/L (14.6 mM) cellobiose. To test for inorganic carbon utilization, 20 mM sodium bicarbonate was added to cell cultures also fed with 5 g/L cellobiose. We used the DSM122-defined medium referred to as CTFUD medium (33). The growth medium was deoxygenated by gassing with argon for 20 min and autoclaved before use. Plasmid and Strain Construction. C. thermocellum DSM1313-derived mutant (∆hpt, also referred to as KJC315) was defined as the wild type and the parental strain to all KO mutants in this study. The gene hpt (clo1313_2927) encodes for hypoxanthine phosphoribosyl transferase, which is used as a counterselection method in the presence of 8-azahypoxanthin. The mutants ∆hpt and ∆hpt∆pflA (KJC318) were constructed using plasmids pKJC83 and pKJC86, respectively. The markerless deletion of the pflA gene (encoded by clo1313_1716) was deleted following the selection and counterselection schemes previously described (1). To delete potential PFOR enzymes (∆hpt∆clo1313_1615-1616, ∆hpt∆clo1313_0673, ∆hpt∆clo1313_0382-0385, ∆hpt∆clo1313_0020-0023, and ∆hpt∆clo1313_1353-1356), plasmids with a configuration similar to pPL161 (Fig. S2A) were constructed. These plasmids designed for gene KO contain a 5′ flanking region and a 3′ flanking region (1 kb each) homologous to the regions upstream and downstream of the target gene(s) to be knocked out in the C. thermocellum genome. Inserted between the 5′ and 3′ flanking regions is a chloramphenicol/thiamphenicol-resistant gene (cat) driven by glyceraldehyde-3-phosphate dehydrogenase (GAPDH) promoter. In addition, thymidine kinase (tdk) from Thermoanaerobacterium saccharolyticum was cloned after the phosphoglycerate kinase promoter for counterselection with 5-fluoro-2′-deoxyuridine. To transform the bacteria with a plasmid, wild type was grown in CTFUD medium at 50 °C anaerobically to OD 600 = 0.8–1. The culture was chilled on ice for 10 min, and cells were collected by centrifugation in a 50-mL Falcon tube at 4 °C and 6,500 × g for 60 min. Supernatants were then removed aerobically. Cell pellets were resuspended with 10 mL of ice-cold 10% glycerol twice, and centrifuged at 4 °C and 6,500 × g for 45 min. Lastly, pellets were resuspended with 100 μL of ice-cold 10% glycerol aerobically. For each transformation, 20 μL of the competent cells was mixed with about 300–600 ng of DNA in 1-mm-gap prechilled electroporation cuvettes (Molecular BioProducts). The mixtures were electroporated (1.2 kV, 1.5 ms, square pulse) aerobically or anaerobically with a BioRad GenePulser XCell (BioRad Laboratories). Cells were immediately resuspended in 1 mL of prewarmed CTFUD medium and rescued at 50 °C anaerobically for 0–3 h. The cultures were then diluted with 25 mL of prewarmed CTFUD medium and incubated at 50 °C anaerobically until the OD 600 was larger than 1. One hundred microliters of growing culture and four 100-fold serial dilutions were then mixed with 25 mL of melting CTFUD agar containing 20 μg/mL thiamphenicol and 10 μg/mL 5-fluoro-2′-deoxyuridine to isolate strains that have undergone double cross-over. The correct genotype was determined by colony PCR and DNA sequencing (Laragen). The loss of the plasmid in all mutants was validated with PCR using primers specific to the plasmid backbone (Table S2), as well as the loss of resistance to thiamphenicol in the markerless deletion of pflA. pH-Controlled Bioreactor Setup. To assess the effect of bicarbonate on C. thermocellum growth, pH-controlled (pH 7.0) fermentation was conducted in batch mode with and without adding 20 mM sodium bicarbonate using bioreactors with a built-in pH controller (Electrolab Biotech and Applikon). Briefly, on the setup of each experiment, 1.8 L of defined CTFUD medium in bioreactors was adjusted to 7.0 and then sterilized by autoclave. Filter-sterilized cellobiose was added to all bioreactors to reach a final concentration of 5 g/L. The bioreactors were deoxygenated by sparging with N 2 gas at 60 cubic centimeter. The N 2 gas flow was turned off, and the bioreactors were sealed once they had become anaerobic. Anaerobic sodium bicarbonate solution was added to the designated bioreactors. The final working volume was adjusted to 2 L for all bioreactors. The same source of inoculum was used for fermentations with and without sodium bicarbonate in parallel in the same model of bioreactors. The pH was monitored and maintained at 7.0 throughout. Quantitative Analysis of Fermentation Products. Cell growth was measured as a function of optical density by spectrophotometry (DU800; Beckman Coulter) at OD 600 . An OD 600 of 1 correlated to 1.04 g/L cell dry weight (R2 = 0.9918). The composition of C. thermocellum biomass was determined to be C 5 H 8 NO 2 by elemental analysis. Briefly, C. thermocellum was grown to stationary phase on 5 g/L cellobiose; the biomass was pelleted and washed three times before drying overnight at 105 °C and subsequently sent to Huffman Labs for analysis of carbon, hydrogen, nitrogen, oxygen, and sulfur. Hydrogen gas was measured using a gas chromatograph (7890A GC; Agilent Technologies) equipped with a thermal conductivity detector and a stainless-steel Supelco 60/80 Mol Sieve column (6 ft × 1/8 in) with argon as the carrier gas. Peak areas were compared with a standard curve, taking into account both the temperature and pressure. Cellobiose, lactate, formate, acetate, and ethanol were measured by HPLC (1200 series; Agilent Technologies) with a mobile phase of 4 mM H 2 SO 4 using an Aminex HPX-87H column with a Micro Guard Cation H Cartridge. The column temperature was set to 55 °C, and the flow rate was 0.6 mL/min. 13C-Tracer Experiment. We adopted a steady-state labeling strategy (2). The 13C-labeling experiment was performed with 20 mM 13C-bicarbonate or 13C-formate, respectively, supplemented into the CTFUD defined medium in addition to 14.6 mM cellobiose. For cell growth, C. thermocellum strains were inoculated into these media with a starting OD 600 of 0.05. When late log-phase was reached (OD 600 ∼ 0.8), 3 mL of cultures was sampled. NMR Analysis. The test sample was collected by filtering growth medium from the wild-type (∆hpt) cells that proliferated with 20 mM 13C-bicarbonate and 14.6 mM unlabeled cellobiose. The reference samples were prepared by dissolving either 13C-formate or 13C-bicarbonate in defined CTFUD medium. All NMR spectra were collected in 5-mm tubes on a 400-MHz (9.4-T) Bruker Avance III HD Nanobay, running Topspin 3.5 with a double-resonance, broad-band observe, 5-mm room temperature probe. One-dimensional 13C spectra were collected at 298.15 K using a 30° flip angle pulse-acquire pulse sequence, a 1.36-s acquisition time, and a 2-s delay. Spectra were collected both with and without 1H decoupling during acquisition to measure J HC . All spectra were referenced to 3-(Trimethylsilyl)propanoic acid (TSP), processed with a shifted sine bell apodization function, zero-filled to 65,000 points using NMRPipe (3), and plotted using Nmrglue (4). GC/MS and Isotopomer Analysis. The sample treatment and GC/MS analysis were done as previously reported (5), with a few modifications. Briefly, 3 mL of sampled cultures was centrifuged at 10,000 × g for 1 min, and the cell pellets were digested with 500 μL of 6 M HCl at 105 °C for 12 h. The hydrolysate was dried under nitrogen gas flow at 65 °C, and dissolved in 50 μL of water-free dimethylformamide. For the GC/MS measurement, the proteinogenic amino acids were derivatized before analysis. The dried hydrolysate, dissolved in N,N-dimethylformamide, was derivatized by TBDMS with 1% tert-butyl-dimethylchlorosilane at 85 °C for 60 min. One microliter of the sample in the organic phase was loaded onto the GC/MS instrument [Agilent GC-6890 gas chromatograph equipped with an Agilent 19091J-413 column (30 m × 0.32 mm × 0.25 μm) directly connected to a MS-5975C mass spectrometer]. Helium was the carrier gas. The oven temperature was initially held at 50 °C for 2 min; it was then raised to 150 °C at 5 °C/min and held at that value for 2 min. Finally, it was raised to 320 °C at 7 °C/min and held at that final value for 2 min. Other settings included splitless and electron impact ionization at 70 eV. The amino acids, including alanine, aspartate, glutamate, glycine, histidine, isoleucine, leucine, methionine, phenylalanine, proline, serine, threonine, tyrosine and valine, were separated and analyzed. To analyze the isotope labeling pattern of amino acids, a mass isotopomer distribution vector, MDV α , was assigned according to Nanchen et al. (16): MDV α = [ ( m 0 ) ( m 1 ) ⋮ ( m n ) ] ∑ i = 0 n m i = 1 , [S1]where m 0 is the fractional abundance of molecules with monoisotopic mass and m i > 0 is the abundance of fragments with heavier masses. The GC/MS data were corrected for the naturally occurring isotopes of oxygen (O), hydrogen (H), and carbon (C) atoms using a correction matrix (Eq. S2) as described by Nanchen et al. (16): MDV α ∗ = C corr , COH - 1 . MDV α , [S2]where MDV α * is the corrected mass isotopomer distribution vector and C corr,COH −1 is the correction matrix. According to Eq. S3, the resulting MDV α * values were then used to assess the fractional labeling enrichment of amino acids, respectively: FL = ∑ i = 0 n i . m i n . ∑ i = 0 n m i , [S3]where n represents the number of carbon atoms in the amino acid and i is the mass isotopomer. Feeding Cells CO 2 /H 2 for Growth. Three methods of feeding C. thermocellum CO 2 /H 2 were used to test its ability to grow entirely on CO 2 with and without H 2 . Briefly, the first method directly inoculated Δhpt (KJC315) cells in two sets of sealed tubes containing CTFUD-defined medium supplemented with 50 mM bicarbonate. One tube was flushed with 100% H 2 gas in the headspace, and the other set of tubes did not have H 2 gas. The initial OD 600 was ∼0.08, and no growth was observed in either set of sealed tubes. In the second method, Δhpt (KJC315) cells were first grown in 5 g/L cellobiose in defined medium to OD ∼ 0.8. Cells were then anaerobically centrifuged at maximum speed for 3 min on a tabletop centrifuge and resuspended in fresh minimal medium with twofold dilution to get to OD 600 ∼ 0.4. Cells were then split into two tubes, and both cultures were supplied with 50 mM 13C-bicarbonate, but one set of tubes was purged with 100% H 2 gas in the headspace and the other without. The third method involved directly feeding the sealed tubes of cultures with a gas mixture of 80% H 2 with balance CO 2 in the headspace after cells were grown up, resuspended in fresh defined medium with twofold dilution to OD 600 ∼ 0.4. OD 600 , CO 2 , and H 2 in the headspace were measured at t 0 and 24 h after t 0 . For both the second and third methods, a tube of cells was inoculated with 5 g/L cellobiose after spinning as a positive control to ensure that cells were viable after centrifugation and to monitor potential growth lag introduced by the intervention.

Acknowledgments We thank Chris Urban, Maria Chun, Sawako Konishi, and Sharon C. Xu for their technical assistance. This work was supported by the National Renewable Energy Laboratory (NREL) Director’s Fellowship (Laboratory Directed Research and Development Subtask 06271403) (to W.X.), and the US Department of Energy (DOE) Energy Efficiency and Renewable Energy (EERE) Fuel Cell Technologies Office (K.J.C., L.M., and P.-C.M.) under Contract DE-AC36-08-GO28308. L.W. and research conducted at the University of California, Los Angeles (P.P.L. and J.C.L.) were supported by the BioEnergy Science Center (BESC), a US DOE Bioenergy Research Center supported by the Office of Biological and Environmental Research in the DOE Office of Science.

Footnotes Author contributions: W.X., P.P.L., L.M., L.W., J.C.L., P.-C.M., and K.J.C. designed research; W.X., P.P.L., L.M., L.W., and K.J.C. performed research; W.X., P.P.L., L.M., L.W., J.C.L., P.-C.M., and K.J.C. analyzed data; and W.X., P.P.L., L.M., L.W., J.C.L., P.-C.M., and K.J.C. wrote the paper.

The authors declare no conflict of interest.

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