Electrosynthesis of Commodity Chemicals by an Autotrophic Microbial Community

Abstract

A microbial community originating from brewery waste produced methane, acetate, and hydrogen when selected on a granular graphite cathode poised at −590 mV versus the standard hydrogen electrode (SHE) with CO₂ as the only carbon source. This is the first report on the simultaneous electrosynthesis of these commodity chemicals and the first description of electroacetogenesis by a microbial community. Deep sequencing of the active community 16S rRNA revealed a dynamic microbial community composed of an invariant Archaea population of Methanobacterium spp. and a shifting Bacteria population. Acetobacterium spp. were the most abundant Bacteria on the cathode when acetogenesis dominated. Methane was generally the dominant product with rates increasing from <1 to 7 mM day⁻¹ (per cathode liquid volume) and was concomitantly produced with acetate and hydrogen. Acetogenesis increased to >4 mM day⁻¹ (accumulated to 28.5 mM over 12 days), and methanogenesis ceased following the addition of 2-bromoethanesulfonic acid. Traces of hydrogen accumulated during initial selection and subsequently accelerated to >11 mM day⁻¹ (versus 0.045 mM day⁻¹ abiotic production). The hypothesis of electrosynthetic biocatalysis occurring at the microbe-electrode interface was supported by a catalytic wave (midpoint potential of −460 mV versus SHE) in cyclic voltammetry scans of the biocathode, the lack of redox active components in the medium, and the generation of comparatively high amounts of products (even after medium exchange). In addition, the volumetric production rates of these three commodity chemicals are marked improvements for electrosynthesis, advancing the process toward economic feasibility.

INTRODUCTION

The U.S. economy is heavily reliant on the use of fossil-based carbon to produce many commodity chemicals and fuels. However, due to supply difficulties, the inevitable decline of these resources, increased world demand, and environmental concerns, a shift away from coal and oil to alternative energy sources such as natural gas, solar, and wind is occurring. However, most of these energy sources are either limited by fluctuations in price and availability or are nonrenewable, as in the case of natural gas. These factors have encouraged research into the development of renewable energy technologies powered by microbes. Of particular interest are microorganisms that can capture the global greenhouse gas CO₂ and convert it to a valuable commodity such as a fuel.

Bioelectrochemical systems (BESs) include microbial fuel cells (MFCs), microbial electrolysis cells (MECs), and electrosynthetic biocathodes (4, 17, 18, 27). Of these, the bioanodes of MFCs and MECs have been the most intensively investigated. The newest and arguably most promising of these technologies is the generation of valuable chemicals by electrosynthesis. Microbial electrosynthesis requires microorganisms to catalyze the reduction of CO₂ by consuming electrons on a cathode in a BES.

The purpose of the present study was to establish a sustainable biocathode from a mixed microbial community that could fix CO₂ and convert it to a commodity chemical. This was achieved by selecting cathodophilic microorganisms from an inoculum obtained from brewery wastewater. Acetate, methane, and eventually hydrogen were the predominant products repeatedly formed at a cathode potential of −590 mV versus the standard hydrogen electrode (SHE). Electrochemical evidence suggests that electron transfer between the electrode and microbes in a biofilm is operating in the absence of soluble redox active components in the medium. This is the first report of the concomitant production of acetate, methane, and hydrogen through electrosynthesis. The production rates of these compounds surpass what has been reported in the literature and was accomplished at a comparatively high cathode potential.

MATERIALS AND METHODS

Source of microorganisms and initial screening.

The biocatalysts described here were enriched from samples taken from a retention basin for brewery wastewater at Palmetto Brewing Company in Charleston, SC. To screen for initial product formation, the brewery wastewater sludge was used to inoculate 20-ml chambers of small BES reactors equipped with graphite rod cathodes. The reactors were poised from −1,000 to −400 mV versus SHE with the goal of selecting for the highest rate of product formation at the highest potential to limit energy input into the system. Products (acetate and methane) were detected after 28 days of incubation at −590 mV and again after the medium had been exchanged once. Controls without voltage applied were monitored for production due to fermentation of the wastewater. Once production free of fermentation was indicated, inocula from these reactors were then transferred to larger three-electrode BES reactors described below in order to further enrich and evaluate the electrosynthetic community.

Three-electrode bioelectrochemical systems.

The BESs consisted of two identical custom designed glass chambers (Chemglass Life Sciences, Vineland, NJ) that had two crimp-seal, butyl rubber sampling ports, a threaded o-ring sealed port for the reference electrode, and a clamped o-ring junction for the membrane (see Fig. S1 in the supplemental material). The total volume of the glass chamber was 150 ml. The two glass chambers were separated by a proton exchange membrane (Nafion 117; The Fuel Cell Store, Boulder, CO) and sealed with an o-ring and clamp. The reference electrode was Ag wire coated with AgCl and immersed in 3 M KCl saturated with AgCl (+210 mV versus SHE). All potentials are reported versus SHE. Both glass chambers contained 30 g (dry weight) of pretreated graphite granules of heterogeneous sizes ca. 10 by 5 by 3 mm and smaller (Showa Denko). Granules were on average 2 g/ml. A 0.9525-cm diameter by 3-cm long pretreated graphite rod current collector connected to a 0.065-cm titanium wire was buried in the graphite granule bed. The graphite electrodes were pretreated by first sonication in deionized water and then washed with acetone, 1 M hydrochloric acid, 1 M sodium hydroxide, and deionized water in succession to remove organic and metal contamination.

The cathode chamber (biotic) was filled with 75 ml of freshwater medium containing per liter: 2.5 g of sodium bicarbonate, 0.6 g of sodium phosphate monohydrate, 0.25 g of ammonium chloride, 0.212 g of magnesium chloride, 0.1 g of potassium chloride, 0.03 g of calcium chloride, 20 ml of vitamin solution, and 20 ml of mineral solution. The vitamin solution contained (per liter): 2 mg of biotin, 2 mg of folic acid, 10 mg of pyridoxine-HCl, 5 mg of thiamine-HCl·2H₂O, 5 mg of riboflavin, 5 mg of nicotinic acid, 5 mg of d-Ca-pantothenate, 0.1 mg of vitamin B₁₂, 5 mg of p-aminobenzoic acid, and 5 mg of lipoic acid. The mineral solution contained (per liter): 1.5 g of nitrilotriacetic acid, 3 g of MgSO₄·7H₂O, 0.5 g of MnSO₄·H₂O, 1 g of NaCl, 0.1 g of FeSO₄·7H₂O, 0.152 g of CoCl₂·6H₂O, 0.1 g of CaCl₂·2H₂O, 0.085 g of ZnCl₂, 0.01 g of CuSO₄·5H₂O, 0.02 g of KAl(SO₄)₂·12H₂O, 0.01 g of H₃BO₃, 0.01 g of NaMoO₄·2H₂O, 0.03 g of NiCl₂·6H₂O, 0.3 mg of Na₂SeO₃·5H₂O, and 0.4 mg of Na₂WO₄·2H₂O. The anode chamber (abiotic) contained a similar medium composition but without the vitamins or minerals and with increased potassium chloride to 1 g/liter and sodium chloride to 2 g/liter. The medium was prepared under anaerobic conditions (80:20 [vol/vol] N₂-CO₂) and passed to the chambers of the BES in an anaerobic glove bag (Coy Laboratory Products). After transfer of the medium, the BESs were removed from the anaerobic chamber, and the headspace was flushed with 80:20 (vol/vol) N₂-CO₂ before inoculation. The BESs were operated in batch mode at 25 ±2°C, and medium exchanges were accomplished by decanting over 90% of the liquid volume, leaving only the granules and what liquid remained in the granular electrode bed. The medium exchanges and subculturing were done in an anaerobic chamber by transferring ∼10 ml of liquid and a small amount (1 to 5 g) of graphite granules from the current-consuming, product-producing reactor into sterile BESs. Where noted, BESs were flushed with 100% CO₂ by using a long needle aseptically pierced through the stopper into the liquid and another short needle in the headspace as gas effluent. To inhibit methanogenic Archaea and enrich for acetogens, 10 mM 2-bromoethanesulfonic acid was added at the time of a medium exchange to a reactor actively producing methane and acetate by electrosynthesis.

Electrochemistry.

During most of the experiments the cathode was poised chronoamperometrically at −590 mV. On day 28 of biocathode operation, the replicate working electrodes (cathodes) shown in Fig. 2 were subjected to cyclic voltammetry (CV). The scan range of the CV was from −200 mV to −1,000 mV, and the scan rate was 1 mV/s. All electrochemistry was performed using a VMP3 potentiostat (Bio-Logic USA). Coulombic efficiencies were calculated by dividing coulombs found in the product (C_p) by total coulombs consumed (C_T). C_p = b·n·F, where b is the number of electrons in the product, n is the number of moles of product, and F is Faraday's constant (96,485 C/mol). C_T was calculated by integrating the area under the current-versus-time curve (i-t curve).

Fig 2.

Replication of biocathodes at −590 mV. (A and B) Coproduction of acetate and methane (A) and coulombs consumed (B) after transfer of the brewery waste biocathode. (C and D) Production of hydrogen (C) and coulombs consumed (D) at −590 mV in abiotic (sterile, uninoculated) control BESs. The BESs were flushed with 100% CO₂ for 30 min on days 7 and 11. Error bars indicate the standard deviations (n = 3).

Analytical methods.

Fatty acids were measured using a high-pressure liquid chromatography (HPLC) apparatus (Shimadzu) equipped with a UV detector at 210 nm. The mobile phase was 0.005 M H₂SO₄ and had a flow rate of 0.55 ml/min through an Aminex HP-87H column (Bio-Rad, Hercules, CA). Methane and hydrogen were measured on a HP6890 gas chromatograph (GC) equipped with an HP-PLOT Molesieve 5A column (30 m by 530 μm by 25 μm) and a thermal conductivity detector. The oven was held at 50°C for 2 min and then increased by 25°C/min to 170°C and held for 0.2 min. The injector temperature was 120°C, and the detector was temperature 250°C. Argon was the carrier gas.

Scanning electron microscopy (SEM).

Graphite granules from the cathode were fixed in 2% glutaraldehyde in 0.1 M sodium cacodylate buffer for 3 h. The granules then underwent a 2.5% osmium tetroxide postfix wash for 1 h. The granules were then dehydrated by a series of ethanol washes (25, 50, 75, 95, and 100%). The samples were sputter coated with gold and palladium with a 100-Å coating (Denton Vacuum). Images were taken with a JEOL JSM-5600LV scanning electron microscope.

RNA extraction.

Samples for RNA extraction were either collected directly into TRIzol (Invitrogen; for MEC granules) or concentrated onto a Sterivex filter (Millipore; PES membrane, 0.22-μm pore size, for MEC supernatant), which was then stored in TRIzol. Samples in TRIzol were incubated at room temperature for at least 15 min and then frozen at −80°C until further processing, as outlined in the supplemental material.

RT-PCR amplification and 16S rRNA sequencing.

Reverse transcription (RT) was carried out with 100 ng of total RNA using random hexamers (SuperScript III; Life Technologies) according to manufacturer's instructions. PCR was performed with either universal bacterial or archaeal primers for the V1-V3 or V2-V3 region of 16S rRNA (see Table S1 in the supplemental material) with the following final concentrations: 1× Green GoTaq reaction buffer, 1 mM MgCl₂, 0.2 mM deoxynucleoside triphosphates, 0.2 μM forward primer mix (equal molar concentrations of degenerate and less-degenerate primer), 0.2 μM reverse primer, 0.625 U of Taq polymerase (Promega), and 0.5 μl of RT reaction per 25-μl PCR volume. Two replicate PCRs were carried out with each of the two following cycling protocols (for a total of four replicates) to maximize priming coverage. The first protocol consisted of an initial denaturing step (94°C, 5 min), 10 amplification steps (45 s each of 94°C, 62°C [decreasing 0.5°C per step], and 72°C), an additional 15 amplification steps (45 s each of 94, 57, and 72°C), followed by a final 10 min extension at 72°C. The second protocol designed to target GC-rich templates (19) is the same as the first, except that all annealing steps were performed for 6 s instead of 45. All PCR replicates were pooled (four total), cleaned (PCR cleanup kit; Qiagen), and quantified (NanoDrop). Amplicons were sequenced on a PacBio-RS sequencer (Engencore, LLC) using a 45-min run time and standard protocols (11). Pacific Biosciences FASTAQ formatted circular consensus sequences have been submitted to the GenBank sequence read archive under SRA056302.

Taxonomic classification.

Sequences were preprocessed and analyzed using Mothur v.1.25 and v.1.27 (29, 30). Briefly, sequences with any of the following features were removed: average quality score of <25, anomalous length (<300 or >615 bp), an ambiguous base (quality score < 1), >8 homopolymers, or >1 mismatch to the barcode or primer. Remaining reads were dereplicated, grouped with similar fragments, and aligned against the Greengenes core database (7) using kmer searching (8mers) with Needleman-Wunsch global, pairwise alignment methods (20). The primers were then trimmed from each read: the B27f primer corresponds to Greengenes alignment positions 109 to 136, A109f corresponds to positions 455 to 493, and U529r corresponds to positions 2232 to 2260. Resulting reads shorter than 300 bp or those likely due to sequence error (14) or chimeras (9) were removed. The reads were then classified using a Bayesian approach and bootstrap cutoff of 80 (34) against the SILVA database (26).

RESULTS

Establishing an autotrophic biocathode.

A three-electrode BES (see Fig. S1 in the supplemental material) was inoculated from a brewery waste culture that was initially screened in a small two-electrode BES. The three-electrode BES was operated for 3 months at a fixed cathode potential of −590 mV. The electrode was the microbial community's only electron donor and CO₂ its only carbon source for growth throughout all experiments. During the first 10 days of incubation, the reactor generated 1.8 mM acetate, followed by 2.6 mM methane, over 30 days as the main products from CO₂ fixation before the first exchange of the spent growth medium (Fig. 1A). Production rates reached 0.18 mmol of acetate per liter of cathode liquid volume per day (mM day⁻¹) and 0.12 mM day⁻¹ methane during this initial startup. Subsequently, after successive medium exchanges, methanogenesis became the dominant process and reached 0.78 mM day⁻¹.

Fig 1.

Development of an electrosynthetic biocathode at −590 mV versus SHE. (A) Operation of a BES for 108 days. Complete replacement of the medium was completed on days 30, 36, 50, 57, and 91. The BES was flushed with 100% CO₂ for 30 min on the days marked with gray arrows. (B) Distribution of coulombs in products compared to total coulombs consumed after the first flushing of CO₂.

As CO₂ was consumed and reduced to methane, the pH in the cathode chamber would frequently exceed 8 (Fig. 1A). To remedy this, 100% CO₂ was flushed through the reactor for 30 min, which then lowered the pH of the medium to ∼6.5. Unexpectedly, this CO₂ flush also revived the production of acetate. The increase in acetogenic activity after CO₂ flushing resulted in rates reaching 1.02 mM day⁻¹ with accumulation of >9 mM in the cathode chamber over 17 days. Methanogenesis also increased in response to the flushing of CO₂, reaching a rate of 1.58 mM day⁻¹. During the 17 days after the start of CO₂ flushing the coulombic efficiency reached 84% (Fig. 1B). To the best of our knowledge, this is the first time the coproduction of acetate and methane has been demonstrated electrosynthetically.

Replication of the autotrophic biocathode.

An important question regarding microbial electrosynthesis resides in the ability to generate sustainable and transferable production rates. After 92 days of operation, supernatant and granules were transferred from the initial reactor into three replicate BESs poised at −590 mV. After a lag period of ∼1 week, product formation began to increase. Once again, acetate and methane were the predominant products in the replicates; however, the acetate production rate was much lower than that of electromethanogenesis (Fig. 2A). Although acetogenesis did not disappear as it did early on in Fig. 1, the rates were not able to compete with methanogenesis, irrespective of the periodic flushing of the cell with 100% CO₂. Over a 10-day period following the initial lag phase, acetate accumulated to 1 mM and methane accumulated to 10 mM. The acetate production rate was 0.1 mM day⁻¹, and the methane production rate reached 1.3 mM day⁻¹ during this period. The coulombic efficiency of the replicates reached 60% (Fig. 2B).

Abiotic (sterile) reactors were also poised at −590 mV to determine whether the abiotic accumulation of hydrogen would be sufficient to account for the methane and acetate observed under biotic conditions (Fig. 2C). At the start of each experiment, ∼0.3 mM hydrogen was immediately produced due to the initial polarization of the cathode. However, from that point forward the abiotic hydrogen production rate was observed at <0.045 mM day⁻¹ over 20 days with a coulombic efficiency ranging from 53 to 64%. Thus, this rate of production cannot account for the mM day⁻¹ rates of methane and acetate production observed in any of the biotic BESs. Coulombs may have been lost in the biotic and abiotic BESs due to gas leakage through joints in the reactor, bubbles trapped in the graphite bed, and in the case of the biotic BESs electrons accumulated into biomass. Despite the portion of electrons unaccounted for, the total coulombs consumed in the biotic replicates far exceeded what was calculated in the abiotic BESs (Fig. 2B and D), indicating microbial catalysis that could not be explained by abiotic hydrogen formation.

Increased rates of electrosynthesis.

The rates of methane or acetate production could be increased by further enrichment of the electrosynthetic biocathodes or by adding a selective inhibitor. After 29 days of operation with repeated medium exchanges (beginning in Fig. 2), the rate of methanogenesis increased, the coproduction of acetate continued, and eventually hydrogen (and occasionally a small amount of formate) was produced (Fig. 3A). The rate of methanogenesis was consistently >1.6 mM day⁻¹ and reached a maximum 7 mM day⁻¹, accumulating to 1.5 mmol in the headspace. The acetate production rate remained near that observed in the initial BES reactor (Fig. 1A), close to 1 mM day⁻¹. Hydrogen did not accumulate to any significant degree until after extended incubation in the experiments documented in Fig. 1 and 2. This was also the case for the experiments presented in Fig. 3 where the microbial community had been further enriched and had experienced multiple medium exchanges. Electrohydrogenesis again lagged behind methanogenesis but suddenly after 7 days of reactor operation increased dramatically to more than 4 mM day⁻¹, eventually reaching 11.8 mM day⁻¹ and accumulated to 1.5 mmol (Fig. 3A). Although hydrogen lagged behind methanogenesis, once it started it was produced concurrently with methane. Also, after an extended lag, formate and acetate eventually were formed at rates of 1 mM day⁻¹ in the methanogenic reactors. The electron recovery (coulombic efficiency) in methane, acetate, formate, and hydrogen was 54% (Fig. 3B). Subsequent transfer cultures in replicate BESs of this community following the establishment of hydrogen production have continued to perform similarly to what is presented in Fig. 3A, generating methane, hydrogen, acetate, and formate.

Fig 3.

Increased rates of electrosynthesis. Two of the replicate BESs described in Fig. 2 were incubated further with two more medium exchanges, the last on day 29. (A) Production of acetate, methane, hydrogen, and formate in one BES maintained without inhibitor. (B) Distribution of coulombs consumed and in all products observed in panel A. (C) Production of acetate and hydrogen in a second BES with 2-bromoethanesulfonic acid added. (D) Distribution of coulombs consumed and in all products observed in panel C. The BESs were flushed with 100% CO₂ on days 33 and 36.

Coproduction of acetate and methane was observed throughout the study (Fig. 1A, 2A, and 3A), but methanogenesis usually outcompeted acetogenesis. This changed upon the addition of the methanogenic inhibitor 2-bromoethanesulfonic acid (Fig. 3C), which resulted in acetogenesis increasing to as high as 4 mM day⁻¹. This rate of activity was sustained in the absence of methanogenesis with subsequent transfers of the treated culture to other BESs. Acetate production started 2 days after medium exchange and inhibitor addition and then increased over the next 10 days, accumulating to 28.5 mM. After a lag of 7 days, hydrogen began to be produced by the community and was then generated concomitantly with acetate (similar to what occurred in the methanogenic reactor in Fig. 3A). The overall rate of hydrogen production was 2 mM day⁻¹ but reached rates of over 9 mM day⁻¹ and accumulated to 1.8 mmol in the headspace. Electron recovery in acetate and hydrogen from the 2-bromoethanesulfonic acid treated community was 67% (Fig. 3D). The biotic production of hydrogen in the reactor with the inhibitor and in the one without (Fig. 3A and C) exceeded abiotic production by at least 200-fold (Fig. 2C and D).

Electrochemical evaluation of the biocatalyst.

Cyclic voltammetry (CV) was performed on the BESs in order to discern possible redox active components associated with the biocathodes. No redox peaks were detected in the abiotic (uninoculated) reactors, indicating a lack of electron shuttles in the medium (Fig. 4, black line). Current production in the abiotic scan was very low at −590 mV and consistent with the low rate of proton reduction observed at this potential over an extended time period (Fig. 2B and D). The CV scan of the abiotic reactor stood in sharp contrast with the catalytic wave seen in the three replicate BESs with live biocathodes producing methane, acetate, and hydrogen (Fig. 4, red line). The onset of catalytic current during the reductive scan of a biocathode was at −340 mV and plateaued at −640 mV versus SHE. The midpoint potential of the catalytic wave was −460 mV, which only varied slightly (approximately ±30 mV) between replicates. The current draw at the peak of the catalytic wave was ∼ 5 mA. In order for the noncatalyzed abiotic BES to reach the same current output, a potential of −900 mV or less was required. The >300-mV discrepancy between peak current in the biotic scan strongly supports microbial catalysis of electrode oxidation.

Fig 4.

Cyclic voltammetry on abiotic (black trace), cell-free supernatant (green trace), and biotic (red trace) BESs. A biotic scan performed on day 28 on replicate reactors shown in Fig. 2. Scan rate, 1 mV/s.

When supernatant (spent media) from the replicate BESs were filtered and inserted into an abiotic reactor, no redox active peaks were observed (Fig. 4, green line). Since no redox active components were observed in the fresh medium or in the filtered supernatant, it is unlikely that a soluble mediator was responsible for electron transfer from the electrode to the microorganisms at −590 mV.

Electrosynthetic microbial community composition.

SEM was used to visualize the prevalence of microorganisms attached to the electrode. Biofilm formation was seen on the graphite granule cathodes from untreated BESs producing acetate and methane (Fig. 5A). The dominant morphology was mostly of rod-shaped microbes varying in size from 2 to 5 μm long. Another, thicker rod shaped organism was also observed. These thicker rods were ∼1 μm long and less prevalent. However, when the biocathode was treated with 2-bromoethanesulfonic acid, these thicker rod-shaped microbes were the dominant morphology on the electrode (Fig. 5B). The observation of these microorganisms on the cathode is consistent with the evidence from the CV, which is supportive of microbial catalyst acting at the surface of the electrode.

Fig 5.

Scanning electron micrographs of electrosynthetic cathode biofilms when primarily methanogenic after 148 days (electrode from the same reactor shown in Fig. 1) (A) and acetogenic after treatment with 2-bromoethanesulfonic acid (day 56, the electrode was from the same reactor shown in Fig. 3C) (B).

To assess the composition of the active microbial population within the electrosynthetic community, total RNA was extracted from samples taken from supernatant or graphite electrode granules at day 91 when acetogenesis was predominant and day 108 when methanogenesis was predominant as shown in Fig. 1A. Overall, in the culture supernatant, the predominant bacterial phyla were Bacteroidetes, Deferribacteres, Firmicutes, Proteobacteria, Spirochaetes, and Synergistetes (Fig. 6). At day 91, when acetogenesis was the predominant activity, members of the Sulfurospirillum genus accounted for 62.3% of the bacterial reads sequenced in the supernatant with another 15.9% belonging to the genus Wolinella. A modest change occurred on day 108, when methanogenesis was the predominant activity, with Sulfurospirillum spp. remaining as the most abundant but decreasing to 36.0%. Members of the genus Wolinella increased to 22.8% and members of the family Spirochaetaceae increased from 9.5 to 24.2%.

Fig 6.

Percent abundance of 16S rRNA for Bacteria (A and B) and Archaea (C and D) from supernatants (s) and graphite cathodes (g) of the active microbial community on days 91 and 108 (yellow arrows in Fig. 1A).

A more dramatic change in the active bacterial population was observed with the samples extracted off the graphite granule electrodes. Acetobacterium spp. were relatively minor members of the supernatant community, but when acetate was the major product (day 91) the percentage of Acetobacterium on the electrode rose to 60.3% (Fig. 6). When methane again dominated and acetate production was low (day 108), the Acetobacterium spp. decreased to 4.7%. An unclassified family (WCHB1-69) from the Sphingobacteriales represented 8.0% of the active population on the electrode at day 91 but became the dominant bacteria at day 108 (37.7%). In contrast, the abundance of WCHB1-69 was relatively constant at approximately 4 to 7% in the supernatant at days 91 and 108. Also found on the cathode on day 91 were members of the family Rhodobacteraceae (8.0%) and the genus Sulfurospirillum (7.4%). Additionally, on day 108, rRNAs of the Synergistaceae family (11.1%) and the Spirochaetaceae family (17.4%) were detected on the cathode.

The predominant archaeal sequences were from the genus Methanobacterium, constituting >93% of the total sequenced archaeal reads, regardless of whether the supernatant or electrodes were examined or when the samples were taken. It is important to note that while acetogenesis was predominant at day 91, methanogenesis was also occurring at both the day 91 and the day 108 time points. Methanobrevibacter represented ∼5% of the reads and unclassified sequences made up a low percentage of total archaeal reads(<1%). See the supplemental material for additional details of the phylogenetic results.

DISCUSSION

An autotrophic microbial community from brewery wastewater was selected on a cathode of a bioelectrochemical system for the production of valuable commodity chemicals. Methane, acetate and hydrogen were all sustainably and reproducibly generated electrosynthetically at a cathode potential of −590 mV versus SHE. Each of these products has been generated with microbial biocathodes, but this is the first study to demonstrate their simultaneous production at rates higher than those reported in the literature. Furthermore, it is the first report of the electrosynthesis of acetate from CO₂ by a mixed microbial community. Differences in laboratory approaches can complicate the comparison of production rates, but sustained rates of methanogenesis and acetogenesis based on cathode volume surpassed what has thus far been discovered for electrosynthesis of these compounds at potentials higher than −700 mV (Table 1).

Table 1.

Comparison of reported rates of electrosynthesis

Product	Cathode potential (mV vs SHE)	Maximum rate (mM day−1)	Microbial source (reference)
Hydrogen	–700	25.3 (3.2 abiotic)	Wastewater (28)
	–900	8.0 (1.5 abiotic)	Desulfovibrio paquesii (2)
	–590	11.8 (0.045 abiotic)	Brewery wastewater (this study)
Methane	–800	1.6	Wastewater (5)
	–800	0.4	Wastewater (33)
	–900	2.1	Wastewater (33)
	–439	0.73	Sediment (25)
	–539	0.54	Sediment (25)
	–590	7.0	Brewery wastewater (this study)
Acetate	–400	0.17a	Sporomusa ovata (22)
	–590	4.0	Brewery wastewater (this study)

^aThis rate is based on the total liquid volume used in a continuous system for 6 days.

A distinguishing feature of the biocathodes examined here was the electrochemical evidence for direct electrode oxidation by the mixed microbial community. Hydrogen production facilitated by the microorganisms may shuttle electrons to the methanogenic and acetogenic microorganisms, but several pieces of evidence indicate that direct electron transfer is also participating: the expression of a catalytic wave observed by CV with an onset at −340 mV and midpoint potential at −460 mV, the lack of similar peaks with sterile or spent media, biofilm formation on the electrode, delayed exponential production of hydrogen, and the recovery of electrons in all three products that exceeds the abiotic generation of hydrogen by several hundred fold.

Electrosynthesis of methane.

Sustainable rates of methane production above 1.5 mM day⁻¹ were achieved and reached 7 mM day⁻¹. Both of these volumetric rates are as high as or greater than any reported in the literature with cathodes poised at potentials above −800 mV (Table 1). Pisciotta et al. recently reported methanogenesis (0.73 mM day⁻¹) at −439 mV that unexpectedly decreased as the potential was lowered to −539 mV, which led the authors to discuss the possibility of organic substrates contributing to the initial rates observed at −439mV (25). Cheng et al. and Villano et al. both demonstrated that lower potentials would support higher methane productivity (5, 33). However, even with increased inputs of energy, the volumetric rates were less than reported here with a cathode potential of −590 mV. There could be numerous reasons for the higher rates observed with the brewery waste electrosynthetic community, including the source of microorganisms, the selection and adaptation of microbes at the chosen cathode potential, and the design and material of the electrode (graphite granules in this case). Regardless, the results presented here clearly indicate that on a working volume basis the rates of methanogenesis far surpass abiotic hydrogen production. Furthermore, we have demonstrated that elevated rates of sustainable methane production may be achieved at potentials above −800 mV.

Electrosynthesis of acetate.

Acetate production concomitant with methane and hydrogen production in the initial BES reached 1.02 mM day⁻¹, a rate that is higher than what has been reported for electroacetogenesis. The first report of electroacetogenesis used pure cultures of Sporomusa ovata to produce 1 mmol of acetate over 6 days (0.17 mM day⁻¹) and trace amounts of 2-oxobutyrate in a continuous flow reactor (22). A second report by Nevin et al. demonstrated electroacetogenesis by several other pure culture acetogens, but none matched the production rate of S. ovata (21).

The rate of electroacetogenesis by the brewery waste community increased to 4 mM day⁻¹ after the addition of 2-bromoethanesulfonic acid, an inhibitor of the methyl reductase of methanogens (12). This rate out paces reported rates for electroacetogenesis by S. ovata by >20-fold. However, Nevin et al. demonstrated electroacetogenesis in a continuous flow system (batch systems were examined in the present study) over 6 days with S. ovata at a cathode potential (−400 mV) substantially higher than what was used in the present study (22). Based on the CV analysis of the brewery waste electrosynthetic community, the onset of the catalytic wave began at approximately −340 mV, indicating that rates of electroacetogenesis by the mixed community could be similar to that of S. ovata at the higher potentials. From a productivity standpoint however, maintenance of the mixed community at −590 mV supports a much higher rate of eletroacetogenesis.

Production of hydrogen and possible mechanisms of electron transfer from the cathode.

With enough driving force, a biocathode will produce hydrogen at rates that exceed abiotic production from an electrode (Table 1). Aulenta et al. observed hydrogen production of 8.0 mM day⁻¹ by a graphite cathode poised at −900 mV and inoculated with Desulfovibrio paquesii, which was ∼5-fold more than what was produced in abiotic controls (2). Sustained activity and growth of the organism with the electrode was not determined. Rozendal et al. demonstrated that hydrogen could be produced with a mixed microbial community in a graphite cathode that was poised at −700 mV (28). Initially, the biocathode produced only methane, presumably hydrogenotrophically due to abiotically produced hydrogen. Bicarbonate was removed from the medium to eliminate methanogenesis, and this resulted in the production of up to 25.3 mM day⁻¹ hydrogen (8-fold greater than abiotic production) and no methane for 1,000 h. The removal of bicarbonate from the medium was not possible for the present study since the goal was the sustained electrosynthesis of organic compounds from CO_2. Similar to what was observed by Rozendal et al. (28), hydrogen did not accumulate during the initial stages of the development of the brewery wastewater community on a biocathode. Surprisingly, however, sustainable and transferable rates of hydrogenesis that were nearly half that reported by Rozendal et al. (Table 1) eventually arose concomitant with the production of methane or acetate while the cathode was poised at −590 mV. Whereas the ratios of biotic to abiotic production ranged from 5 to 8 in the previous studies (2, 28), here with the cathode poised at a higher potential the ratio increased to >250 with several hundredfold more electron equivalents simultaneously recovered in methane or acetate.

It is possible that electrons are being directly delivered from the cathode to the microorganisms producing methane, acetate, and hydrogen. It is also plausible that hydrogen could be serving as the electron-carrying intermediate between the electrode and the methanogens and acetogens, but it is evident that such hydrogen must be produced biotically at the cathode. It is clear that the biology of the system is greatly facilitating the electrosynthetic process since the electron recovery in products is so high versus what is recovered abiotically. The catalytic wave detected by CV (Fig. 4), combined with the observation of a biofilm on the cathode and the delayed production of hydrogen concomitant with methanogenesis and acetogenesis, is in agreement with the biological production of hydrogen being coupled to direct electron transfer from the electrode to a microbe. The onset of current draw began at −340 mV (Fig. 4), a cathode potential that was >300 mV higher than the onset of current draw in the abiotic reactors, indicating that the microorganisms catalyzed electron transfer from the electrode. Importantly, the plateau in current is a unique signature of microbial catalysis of electron transfer from the electrode because abiotic current draw would be continuous with decreasing potentials. If the catalytic wave is expressed by proton-reducing bacteria, then the constant supply of electrons from the cathode in a proton-rich environment may enable these microbes to extract energy in the form of ATP while generating hydrogen. Although growth was not measured here, the evidence of a biofilm and sustained and transferable activity suggests that growth did occur. It is conceivable that a syntrophic relationship between electrode-oxidizing proton reducers and acetogens and methanogens may help support the growth of the entire community and result in faster production rates of all three products. Interestingly, however, methane and acetate continue to be produced at fast rates even as hydrogen accumulation increases, indicating that hydrogen does not shutdown proton reduction under these conditions. This is in agreement with what Aulenta et al. observed with D. paquesii producing hydrogen in an electrochemical cell (2). Therefore, either the methanogens and acetogens are unable to keep up with the microbes responsible for hydrogen generation, or perhaps they do not use the free hydrogen and directly receive electrons from the electrode, possibly by direct electron transfer between species (32) or through an electron-carrying mediator other than H₂. However, the lack of any redox peaks in the CV scan of the spent medium would suggest that the medium or the microbial community does not supply a soluble mediator other than hydrogen.

Electrosynthetic microbial community.

Microbial communities are notorious for the intricate interactions between microorganisms that frequently result in an efficient and productive process. This is due to the natural selection of microorganisms that will operate in stable consortia. Often it is desirable to select for such consortia to perform useful reactions, e.g., the synthesis of commodity chemicals, particularly when the growth and survival of the microbial community is dependent on these reactions. Extended incubation in a BES with a poised potential and only CO₂ as the carbon source served as the selection process for this study. When a potential of −590 mV was applied, the result was a community that would electrosynthesize three commodity chemicals: methane, acetate, and hydrogen. A diverse group of active microorganisms were detected on the cathodes with the bacterial community shifting concomitantly with changes in the prevailing functional activity (acetogenesis, methanogenesis, and hydrogenesis).

The data indicate that at least one member of the community will interact directly with the electrode. Acetobacterium spp. were the most prevalent and active Bacteria on the electrode when acetate was produced. Previous attempts to electrosynthesize acetate with Acetobacterium woodii failed, although it consumed H₂ supplied to the cathode chamber (21). The Acetobacterium spp. detected here were strongly associated with the electrode and dominated that population (60.3%). Either these Acetobacterium spp. are quite different from A. woodii or the microbial community on the electrode affords Acetobacterium with advantages unrecognized in the pure culture. The Sphingobacteriales that became dominant as the community progressed have close sequence identities to microorganisms found in electrode reducing biofilms and to hydrogen-producing communities. It is possible that microorganisms such as the Sphingobacteriales WCHB1 or Sulfurospirillum are oxidizing the electrode and generating hydrogen (similar to D. paquesii) that feeds the methanogens and acetogens; however, this could not be proven at this time. Hydrogenotrophic methanogens, Methanobacterium in particular (93%), dominated the Archaea detected on the electrode regardless of conditions, and the dominant microbial morphology observed on the electrode when methanogenic was a rod with the appearance of Methanobacterium. Cheng et al. (5) reported a similar percentage of Methanobacterium in an electromethanogenic cathode. All three dominant members of the varying community discussed above could potentially be responsible for electrode oxidation.

Implications for commodity chemical production.

Methane is the primary component of natural gas (NG), which is widely used in automobiles and electricity generation (3, 10). It is also the primary source of hydrogen for the production of nitrogen fertilizers (1). No biofuel, including electrofuels at this time, could compete economically with the present low price of NG unless subsidized, but the cost of NG will rise as its use increases. In addition, even though a 100-year supply of NG has been estimated (13), it will eventually be consumed. Although it is by far the cleanest of the fossil fuels, its use still results in the release of climate-changing CO₂. Furthermore, the hydraulic fracturing process needed to extract shale gas requires large amounts of water and risks groundwater contamination (23). Electromethane from renewable and sustainable sources of energy will have many of the same benefits but none of these problems, and it could be developed first to supplement NG with the goal of one day replacing it. As the present study helps to demonstrate, the rates of electromethanogenesis can be improved. At 131 mol of methane per gallon of gasoline equivalent (GGE) (based on 114,000 Btu per gallon of gasoline, 1,011 Btu per cubic foot of CH₄, and ideal gas law at 25°C), the 7 mM day⁻¹ rate observed for electromethanogenesis would calculate to a 0.05 GGE day⁻¹ m⁻³ reactor. Although still requiring improvement, increasing this rate by an order of magnitude would conceivably produce 0.5 GGE each day from a reactor the size of a kitchen appliance. As this technology attracts more attention, rates may increase so that a renewable biogas technology to replace NG may be developed.

Acetic acid is another valuable commodity chemical made from fossil fuels that is used in industrial processes to produce vinyl acetate for paints and adhesives and to a smaller extent vinegar (6). Production for human consumption, e.g., food and cosmetics, requires a higher degree of purity, which is achieved by microbial fermentation (8, 24). Acetate is also a key intermediate in the production of biofuels, since it has been shown to be a feedstock for a microbial community to produce ethanol in BESs using methyl viologen as an electron carrier (31). Any biosynthetic pathway that involves reducing CO₂ to multicarbon compounds must first pass through acetyl coenzyme A (acetyl-CoA) and acetate can be readily converted to acetyl-CoA by microbes. Hence, electroacetate could be used as a precursor for fuel production or for the production of high-purity foods and cosmetics. In addition, a synthetic biology approach could be coupled with electroacetogenesis to produce commodity chemicals. A similar approach was taken by Li et al. with formic acid as a feedstock to make isobutanol (15).

Electrosynthesis potentially offers a revolutionary way of producing the chemicals needed to sustain our modern culture. The carbon source for the process, CO₂, is plentiful and inexpensive, the electrons may be supplied from sustainable non-carbon-based sources, and the land mass requirements are negligible and will not compete with food crop production; moreover, strictly carbon neutral electrosynthesis presents an attractive way to combat climate change. Analogous to the field of microbial fuel cells where intensive research has led to a better understanding of the process and exponential gains in current generation (16); here, it has been demonstrated that the rates of production of multiple commodity chemicals by electrosynthesis can be further increased, thereby advancing the technology closer to becoming competitive with the fossil-carbon based industries.