Enhanced Electricity Production by Use of Reconstituted Artificial

Feb 21, 2012 - ABSTRACT: Microbial fuel cells (MFCs) can convert organic compounds directly ... −2 . Power production was by direct transfer of elec...
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Enhanced Electricity Production by Use of Reconstituted Artificial Consortia of Estuarine Bacteria Grown as Biofilms Jinwei Zhang,†,∥,⊥ Enren Zhang,‡,∥ Keith Scott,§ and J. Grant Burgess*,† †

School of Marine Science and Technology and Centre for Bacterial Cell Biology and §School of Chemical Engineering and Advanced Materials, Newcastle University, Newcastle NE1 7RU, United Kingdom ‡ Department of Chemistry and Chemical Engineering, Yangzhou University, Yangzhou City 225002, China S Supporting Information *

ABSTRACT: Microbial fuel cells (MFCs) can convert organic compounds directly into electricity by catalytic oxidation, and although MFCs have attracted considerable interest, there is little information on the electricity-generating potential of artificial bacterial biofilms. We have used acetate-fed MFCs inoculated with sediment, with twochamber bottles and carbon cloth electrodes to deliver a maximum power output of ∼175 mW·m−2 and a stable power output of ∼105 mW·m−2. Power production was by direct transfer of electrons to the anode from bacterial consortia growing on the anode, as confirmed by cyclic voltammetry (CV) and scanning electron microscopy (SEM). Twenty different species (74 strains) of bacteria were isolated from the consortium under anaerobic conditions and cultured in the laboratory, of which 34% were found to be exoelectrogens in single-species studies. Exoelectrogenesis by members of the genera Vibrio, Enterobacter, and Citrobacter and by Bacillus stratosphericus was confirmed, by use of culture-based methods, for the first time. An MFC with a natural bacterial consortium showed higher power densities than those obtained with single strains. In addition, the maximum power output could be further increased to ∼200 mW·m−2 when an artificial consortium consisting of the best 25 exoelectrogenic isolates was used, demonstrating the potential for increased performance and underlying the importance of artificial biofilms for increasing power output.

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INTRODUCTION Microbial fuel cells (MFCs) can convert biodegradable and reduced compounds, such as glucose, acetate, and lactate, directly into electricity, offering a clean and renewable source of energy.1−6 In addition, MFCs may also assist environmental protection, for example, through wastewater treatment. In MFC devices, bacterial cells in the anode chamber play a key role in catalyzing the oxidation of an organic substrate (the fuel) and transferring electrons derived from metabolic processes to the electrode.7 Thus, one primary aim of MFC research is to isolate electrogenic bacteria or communities with high electrochemical activity. Electricity can be produced by naturally existing consortia of bacteria without addition of exogenous mediators in fuel cell systems.8,9 Sufficient current can also be generated in order to power subsurface devices by placing an anode into anoxic sediment and a cathode into overlying water.10 The diversity of bacteria capable of exoelectrogenic activity in anodophilic biofilms has been well-studied by culture-independent techniques,8,11 and some pure cultures that exhibit strong electrogenic activity in the MFC environment have been characterized. 7 There is a broad range of confirmed exoelectrogenic bacteria, including the five classes of Proteobacteria as well as the Firmicutes and Acidobacteria.12,13 However, community analysis of electrochemically active biofilms in MFCs suggests a far greater diversity of © 2012 American Chemical Society

exoelectrogens than was previously suspected. Novel electrogenic bacteria remain to be discovered, and importantly, many strains that have been reported from 16S rDNA studies remain to be cultured in the laboratory and have their contribution to electricity production confirmed. The use of biofilm communities may also allow production of greater power densities than individual strains. We were interested in using artificially reconstituted consortia of exoelectrogens to enhance electricity production. Such an approach, where strains showing optimum properties are artificially selected over strains showing poor performance, has been used successfully before, for example, to enhance oil biodegradation.14 In the present study, highly electrogenic anodophilic biofilms from estuarine sediments are reported, and 74 bacterial strains are isolated and their electricity generating properties are surveyed. Our results expand the known diversity of bacteria capable of electricity generation. Furthermore, we could enhance electricity production by deliberately choosing and reconstituting biofilms containing the best electricity producers. Received: Revised: Accepted: Published: 2984

February 13, 2011 January 22, 2012 January 31, 2012 February 21, 2012 dx.doi.org/10.1021/es2020007 | Environ. Sci. Technol. 2012, 46, 2984−2992

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Figure 1. Voltage output produced by (A) sedimentary bacteria-activated MFC and (B) anodophilic bacteria-activated MFC. (■) Voltage output by control MFCs without bacteria. Arrows in panel B indicate the replacement of anodic solution by fresh growth medium. (C, D) SEM images of acetate-induced biofilm morphology on the anode surface. Arrows indicate pilus-like appendages connecting bacterial cells. Biofilms on the anode surfaces were examined by SEM as described previously.16.

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MATERIALS AND METHODS Sediment Samples and Growth Media. Estuarine sediments were collected at low tide from the River Wear (54°54′25″ N, 1°21′35″ W) and stored in an anaerobic GasPak EZ (BD). Bacteria were anaerobically cultured in fumaric acid medium (FAM):15 10 mM fumaric acid, 10 mM sodium acetate, and 0.05% yeast extract in a 1:1 (v:v) mixture of seawater and distilled water, adjusted to pH 7.0, flushed with N2 to remove oxygen, and autoclaved. Sediment samples were inoculated into FAM for 2 weeks, and then 10 mL of the resulting mixed culture was used to inoculate the sterile anaerobic chamber of the MFC. Fresh seawater was collected from the Dove Marine Laboratory, Cullercoats, Tyne and Wear, U.K., and passed through a 0.2 μm pore-size filter for medium preparation. Microbial Fuel Cell Construction and Operation. The dual-chamber MFC was constructed from two glass 250 mL bottles (Corning Inc.) with H2315 carbon cloth (4 × 5 cm) (Freudenberg FCCT KG) electrodes. The MFC chambers were connected by a tubular duct of 1.3 cm inner diameter with a Nafion membrane (Nafion 117, Dupont Co., Wilmington, DE), installed as a separator as described previously.16 The membrane was equilibrated by incubation in 0.1 M NaCl solution for 2 h prior to use. New electrodes were pretreated by soaking in 1 M HCl to eliminate possible metal ion contamination. Before inoculation, the anode chamber was filled with 200 mL of medium, containing 10 mM sodium acetate and 0.05% yeast extract in 1:1 diluted seawater, and flushed with pure N2 gas for 20 min to flush out oxygen. The

cathode chamber was filled with 200 mL of electrolyte solution containing 50 mM K3Fe(CN)6 and 100 mM KH2PO4 (pH adjusted to 7.0 with 1 N NaOH). The recovered electrode, coated with biofilm, was used to provide inocula for new microbial fuel cells with fresh medium. In order to study the exoelectrogens and nonexoelectrogens present in anode biofilm, each pure strain and/or combination culture of pure strains was further characterized with the same fuel cell system. All experiments were conducted at room temperature (23 ± 2 °C), and one setup with no inoculum was also operated in parallel as a control. Electrochemical Measurements. MFCs were operated in batch mode and the circuit was operated with a fixed external resistance of 1000 Ω. The voltage across the known resistance was continuously measured by use of a digital multimeter (SkyTronic). Polarization curves for MFCs were measured by linear sweep voltammetry (scan rate 1 mV·s−1) from the open circuit potential to 0 V with a potentiostat (Autolab PGSTAT302) when a stable voltage production was achieved. Power and maximum power were calculated by use of data from stable voltage production and measured polarization curves, respectively, and then normalized to the total area (40 cm2) of the anode surface. Internal resistance was determined by electrochemical impedance spectroscopy. To characterize anodophilic biofilms in situ, cyclic voltammetry measurements were carried out with Autolab PGSTAT302,16 with the anode in the anodic chamber as the working electrode, a Ag/AgCl electrode connected to anodic solution through a Luggin capillary as reference electrode, and a 30 × 10 × 0.2 mm 2985

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Figure 2. (A) Cyclic voltammetry measurement (scan rate 50 mV·s−1) with a glassy carbon electrode on fresh growth medium, anodic supernatant, and anodic suspension during stable electricity generation. (B) Current density−voltage and current density−power density relationships for acetatefed microbial fuel cells during stable electricity generation (measured by slow scan cyclic voltammetry, scan rate 1 mV·s−1).

hours. The medium replacement and voltage restoration of the present MFCs could be repeated without decay in electricity generation. The restoration of voltage after replacement of growth medium suggested that anodophilic biofilms could directly transfer electrons derived from acetate metabolism to the anode surface in the absence of any soluble electron shuttles. This characteristic was also supported by cyclic voltammetry measurements. Figure 2A shows cyclic voltammograms with a glassy carbon electrode for fresh sterile medium without inoculum, MFC-derived cell suspension, and cell-free suspension. No obvious electroactive species were detected in the above solutions within the electrode potential range where possible soluble electron shuttles would be detectable as redox peaks.18,19 The presence and appearance of anodophilic biofilms was examined by SEM, which confirmed coverage by biofilms consisting of different coccoid and rod-shaped bacterial cells. Pilus-like appendages connecting the cells were also observed (Figure 1 C,D). Performance of Microbial Fuel Cells. The open-circuit potentials of the double-bottle MFCs reached 0.8 V when sufficient quantities of acetate were present. When the current became stable for acetate-fed MFCs (voltage output 0.65−0.7 V), slow cyclic voltammetry was carried out to determine the polarization curves of the MFC. Figure 2B shows a representative measurement. The voltage fell almost linearly with increasing current density, which indicates that ohmic resistance was predominant in the MFCs, and no obvious voltage drop caused by charge transfer resistance was observed at a lower rate of current flow. The voltages achieved in the MFC tests agree with values obtained during the biofilm development stage (Figure 1), where a voltage of around 0.65− 0.7 V occurred at current densities around 162−175 mA·m−2. Electrochemical impedance spectroscopy (EIS) (Supporting Information, Figure S1) showed that the internal resistance (IR) of the MFCs used in this work was in the range of 120− 145 Ω, greater than higher specification devices reported previously.20 From the voltage−current density behavior of the MFC (Figure 2) the internal resistance was in the range of 120−145 Ω; these figures are also supported by the polarization plot in Figure 2B and the EIS data (Supporting Information, Figure S1). At low cell voltages there was an indication of a mass transport limitation, which may be associated with the

platinum foil counterelectrode in the cathode chamber. Other electrochemical measurements were conducted in 50 mL threeelectrode cells with a 3 mm diameter glassy carbon electrode as the working electrode, a Ag/AgCl electrode as reference electrode, and a Pt wire as the counterelectrode. Strain Isolation and 16S rDNA Analysis. Samples were inoculated onto different marine isolation media (MA1−MA3): MA1, Difco Marine agar; MA2 (fumaric acid medium, FAM), 10 mM sodium fumarate, 10 mM sodium acetate, 0.05% yeast extract, and 1% agar, made up with 50% by volume of seawater and 50% distilled water; and MA3 (ferric citrate medium, FCM), 5 mM ferric citrate, 10 mM glucose, 0.05% yeast extract, and 1% agar, also with 50% seawater and 50% distilled water. Bacteria colonizing the electrode surface were removed from the biofilm, resuspended in sterile growth medium, spread onto agar plates, and grown under anaerobic conditions by use of the BD Gaspak system (see above) until single colonies were obtained; this was repeated until pure isolates were obtained. The 16S rRNA gene sequences were checked for similarities to DNA sequences in the NCBI and RDPII databases by use of BLAST. The alignment and phylogenetic analysis of sequences were achieved with DNAMAN software package (version 5.2.2), cluster and molecular evolutionary genetics analysis (MEGA) version 2.1, as described previously.17 The nucleotide sequences of 16S rDNA have been deposited in GenBank under accession numbers FN997605 through FN997642. Scanning Electron Microscopy. Biofilms on the anode surfaces were examined by scanning electron microscopy (SEM) on a Cambridge Stereoscan 240 according to established techniques.16

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RESULTS Power Generation. Generally, a long lag phase (over 10 days) occurred before voltage started to increase. Figure 1A shows the voltage output of a fuel cell inoculated with sediment bacteria, with a lag phase of ∼13 days. The voltage began to increase exponentially and reached higher stable electricity generation (>0.6 V), which was maintained for 10 days. The recovered electrode coated with biofilm showed a high stable level of voltage (>0.6 V) after a much shorter lag phase ( 0.65 V). A mature biofilm on the anode was necessary for high stable electricity generation, consistent with the observations on growth medium replacement (Figure 1). When the acetate in the anodic growth medium was depleted from 5 mM, the limiting current of the catalytic wave also decreased (Figure 3B). Phylogenetic Analysis of Anodophilic Bacteria. From the electrode surface, 74 isolates were obtained and their 16S

oxygen reduction reaction or possibly acetate mass transport limitation. The acetate-fed MFC used in this study could generate a maximum power density of 175 mW·m−2 (Figure 2B). Maximum power density and stable power generation were higher than marine and freshwater sediment-inoculated MFCs reported recently (Table 1). This could be attributed to the use of added acetate as a feedstock and also the use of anodophilic microbial biofilms in transporting electrons to the electrode, considering the high internal resistance of the MFCs. In fact the IR and peak power densities were about 130 Ω and 175 mW·m−2, respectively, indicating the potential of relatively high power generation with higher specification MFCs using minimal electrode separation and improved cathodes. It is important to point out, however, that direct comparisons are difficult due to the different laboratory setups and also the high variability in the microbiology of each system. Electrochemical Properties of Anodophilic Biofilms. Cyclic voltammetry (CV) was performed to characterize the catalytic properties of the anodophilic biofilm on the carbon 2987

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Table 2. Anodophilic Bacterial Isolates and Their Power Generation Abilities When Tested in a Single-Species MFC phylogenetic group

a

representative isolates

Bacillaceae Bacillaceae Bacillaceae Enterococcaceae

MS10a MS28; MS27 MS34; MS4; MS1 MS23; MS7; MS51; MS53; MS60; MS69

Rhodobacteraceae

MS22; MS9; MS45; MS46; MS65

Alcaligenaceae Alcaligenaceae

MS11; MS13; MS14; MS50; MS62 MS16; MS47; MS48; MS52; MS61

Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Aeromonadaceae

MS2 MS6 MS25 MS5 MS8 MS17 MS18 MS36; MS39 TRS1-B4 TRS1-WB; MS31; MS49; MS59 MS24; MS20; MS21; MS58; MS66

Aeromonadaceae

TRS1-B1; MS15; MS19; MS33; MS70

Aeromonadaceae

MS12

Vibrionaceae Shewanellaceae Shewanellaceae Moraxellaceae Alteromonadaceae Alteromonadaceae

MS26 MS32 MS43 TRS1-R2; MS42; MS56; MS64; MS67 MS3; MS37; MS54; MS55; MS63 MS38; MS40; MS44; MS57; MS68

Campylobacteraceae

MS30; MS35

Bacteroidetes

TRS1-A1

nearest type strain in GenBank (accession no.) Firmicutes B. stratosphericus 41KF2aT (AJ831841) B. altitudinis 41KF2bT (AJ831842) E. mexicanum 8NT (AM072764) V. fluvialis CCUG 32704T (Y18098) α-Proteobacteria R. maris JA276T (AM745438) β-Proteobacteria A. faecalis subsp. faecalis IAM12369T (D88008) A. faecalis subsp. parafaecalis GT (AJ242986) γ-Proteobacteria K. oxytoca JCM 1665T (AB004754) K. oxytoca JCM 1665T (AB004754) K. oxytoca JCM 1665T (AB004754) E. aerogenes NCTC 10006T (AJ251468) C. freundii DSM 30039T (AJ233408) C. freundii DSM 30039T (AJ233408) C. freundii DSM 30039T (AJ233408) C. freundii DSM 30039T (AJ233408) C. freundii DSM 30039T (AJ233408) S. proteamaculans DSM 4543T (AJ233434) A. salmonicida subsp. masoucida ACC 27013T (X74680) A. hydrophila subsp. hydrophila ATCC 7966T (CP000462) A. hydrophila subsp. hydrophila ATCC 7966T (CP000462) V. azureus LC2−005T (AB428897) S. haliotis DW01T (EF178282) S. algae ATCC 51192T (AF005249) P. nivimaris 88/2−7T (AJ313425) M. lipolyticus SM19T (AY147906) M. lipolyticus SM19T (AY147906) ε-Proteobacteria A. nitrofigilis CCUG 15893T (L14627) CFB Group Bacteria M. fragile JC2469T (FJ394546)

similarity, %

max power density, mW·m−2

99.859 99.930 99.636 99.853

87.5 (±1.3) 6 (±0.7) 0 0

96.726

0

99.430 99.571

0 0

98.517 98.593 98.742 98.655 99.514 99.789 99.220 99.507 99.578 99.647 99.956

9 (±1) 12.5 (±1.7) 5 (±0.6) 15 (±0.7) 38 (±0.7) 30 (±0.5) 10 (±0.6) 14 (±0.5) 9 (±1) 0 0

99.719

15 (±1)

99.859

13.5 (±1.5)

99.127 99.788 99.531 99.929 98.379 98.310

30 (±1) 5 (±0.2) 15 (±0.6) 0 0 0

95.345

8.7 (±0.4)

93.309

12.5 (±0.5)

Strains chosen for phylogenetic analysis are shown in boldface type.

Exiguobacterium mexicanum. Within the Enterococcaceae, one strain similar to Vagococcus fluvialis was recovered. The remaining strains fell into α-Proteobacteria (6.8%), βProteobacteria (10.8%), ε-Proteobacteria (2.7%), and Cytophaga−Flexibacteria−Bacteroides group (CFB) (1.4%). Most of the strains isolated were found to have high homology (98.3−99.9%) to their closest neighbors, while MS22, MS30, and TRS1-A1 showed lower similarities of 96.7%, 95.3%, and 93.3%, respectively, to Rhodobacter maris, Arcobacter nitrofigilis, and Marinifilum fragile. These strains are taxonomically unusual and worthy of more detailed characterization. Electrochemical Properties of the Isolated Anodophilic Bacteria. Of the 44 γ-Proteobacteria isolates, 19 strains could produce electricity. Strains MS8 and MS17, similar to C. freundii, and strain MS26, similar to V. azureus, produced the highest power density of 30−40 mW·m−2 (Table 2). Three out of 15 strains in the Firmicutes group were able to produce electricity. Strain MS10, similar to B. stratosphericus, was found with 87.5 mW·m−2 power density production under the above-

rDNA was sequenced. Thirty-eight distinct sequences were chosen for phylogenetic analysis (Table 2 and Figure 4). NCBI nucleotide BLAST searches using the partial 16S rDNA sequences of these 74 strains revealed that 44 (59.5%) of the isolates fell into the γ-Proteobacteria group and shared a phylogenetic affiliation with members of Enterobacteriaceae, Aeromonadaceae, Vibrionaceae, Shewanellaceae, Moraxellaceae, and Alteromonadaceae. These results were further confirmed by Gram staining and microscopy. Twenty-two strains fell into the class Enterobacteriaceae and showed similarity to four taxonomic units, Klebsiella oxytoca, Enterobacter aerogenes, Citrobacter freundii, and Serratia proteamaculans; six strains fell into the class Aeromonadaceae, including two taxonomic units, Aeromonas salmonicida and Aeromonas hydrophila; while Shewanellaceae contained Shewanella haliotis and Shewanella algae. Isolates similar to Vibrio azureus, Psychrobacter nivimaris, and Marinobacter lipolyticus were also found. The Firmicutes represented 16.2% of the sequences with strains similar to Bacillus stratosphericus, Bacillus altitudinis, and 2988

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Figure 4. Neighbor-joining distance tree based on the nearly complete and aligned 16S rDNA sequences of 38 representative strains observed in this study and their nearest neighbors. The phylogenetic tree was constructed by the neighbor-joining method using the programs of the MEGA package. Bootstrap analysis (1000 trials) was used to provide confidence estimates for phylogenetic tree topologies. Bar = 0.05 nucleotides substitution per site.

mentioned conditions, while B. altitudinis-similar strains MS27 and MS28 generated lower power of ∼6 mW·m−2. In the α-, β-, and ε-Proteobacteria and CFB group bacterial phyla, strain MS11, with homology to Alcaligenes faecalis, was unable to produce electricity, whereas strain MS30, with homology to A. nitrofigilis, and strain TRS1-A1, tentatively identified as M. fragile, showed relatively small power generation (3.5−17.5 mW·m−2). Electrogenic Properties of Artificial Consortia. When 74 strains were remixed and cultured as an artificial biofilm consortium (TRS2) in the MFCs, power output remained similar to that of the wild-type consortium (TRS1). However, the maximum power output was increased to ∼200 mW·m−2

when the consortium biofilm (TRS3) was reconstituted with only the exoelectrogenic strains (Table 2 and Figure 5).

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DISCUSSION

Using a two-chamber microbial fuel cell inoculated with estuarine sediments, we isolated anodophilic biofilms that selectively proliferated in the presence of acetate on carbon cloth electrodes. Our approach involved allowing the microbial community to adapt to this environment, and as a result a power density of 175 mW·m−2 was achieved. In 2001, Reimers et al.10 first utilized marine sediment−seawater interfaces in situ and obtained power generation of 10 mW·m−2 by use of 2989

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detected in a cysteine-fed MFC by use of DGGE, strains of Vibrio were not isolated or independently shown to produce electricity.31 Citrobacter was reported in a bacterial consortium in a glucose/glutamate-fed MFC on the basis of RFLP analysis,38 and Enterobacter was first reported as part of a power-generating consortium digesting cellulose.39 The genus Klebsiella actively produces electricity in MFCs40,41 and in this study (strains MS2, MS6, and MS25). Although in this work acetate is the main electron donor, trace amounts of fermentable substrates from yeast extract used in the medium may also account for nonexoelectrogens present in the anode biofilm.42,43 A significant proportion of nonexoelectrogenic bacteria could be cultured from the original biofilms (Table 2), and it has been suggested that they play a role as helper strains, fermenting trace amounts of organic matter.44 Interestingly, maximum power output was increased and maintained at ∼200 mW·m−2 when artificial consortia were reconstituted with only exoelectrogenic bacteria (Table 2). It may be that the presence of nonexoelectrogenic bacteria can disrupt or reduce the overall electrical conductivity of the biofilm. No detectable redox species were involved in electron transfer by the anodophilic biofilms, and isolates similar to B. stratosphericus, C. freundii, and V. azureus isolated from the biofilms could generate considerable power in the same MFC devices using acetate as electron donor. Therefore, power production could be confirmed by direct transfer of electrons to the anode by the bacterial consortia growing on the anode as demonstrated by cyclic voltammetry (CV). The potential wave observed with the anodophilic bacterial biofilm was similar to results obtained with carbon-attached biofilms formed from a pure strain of Geobacter sulfurreducens.21 G. sulfurreducens may directly transfer electrons from bacterial cells to the anode via several different redox-active outer-membrane cytochromes.21,22 It is unknown which microbial species or proteins in the present complex biofilm play these roles. However, the redox potentials observed (−200 to 0 mV versus SHE) are similar to those of strains or purified outer-membrane cytochromes.21,23,24 Strains of Geobacter were not isolated in this study; however, it is worth noting that Shewanella outer membrane cytochromes also have relatively low redox potentials. Scanning electron microscopy of the biofilms showed numerous pilus-like appendages, connecting cells to form an integrated community (thick biofilm) on the surface of the electrode. Further work is required to determine the precise role of such appendages in electron transfer.45 It will be interesting to examine how bacteria cooperate with each other to efficiently metabolize organic carbon sources for electricity production and to investigate whether the lateral appendages produced by strains isolated in this work contribute to electron transfer. Models of consortium biofilm structure and its activities and a better understanding of the synergistic cooperation of each individual strain in the biofilm will help us to better predict the maximum power densities achievable by MFCs.

Figure 5. Voltage output produced by MFCs containing wild-type biofilm (TRS1, ○), an artificial consortium of pure strains isolated from the original biofilm (TRS2, △), and an artificial consortium composed of all exoelectrogenic bacteria (TRS3, ◇).

graphite fiber-based electrodes. By using different marine sediments and specific anodes, a range of power densities (20−100 mW·m−2) was obtained.25−28 By using high surface area and semienclosed anodes, respectively, the power generation was improved to 14029 and 233 mW·m−2.30 However, in the laboratory, electricity production from sedimentary MFCs remains low, between 40 and 70 mW·m−2.16,31 Much analysis of the composition of bacterial communities inhabiting fuel cell anode chambers has been carried out by culture-independent methods such as denaturing gradient gel electrophoresis (DGGE) or restriction fragment length polymorphism (RFLP) of the amplified 16S rDNA fragments and sequencing of the dominant bands.8,13,25,32,33 However, such analysis provides no information about whether these species are electrogenic. A culture-dependent study, though more difficult, allows relative contributions to the overall power generation of the various strains to be estimated. This is important, since our work demonstrates that a significant proportion (67%) of bacteria isolated from an electrogenic biofilm are not electrogenic when tested in isolation. On the other hand, a limitation of the culture-dependent approach is that only some of the anodophilic bacteria can be cultured. So a combination of these approaches must be used. Cytophaga/Flexibacter/Bacteroides, one of the major groups of the MFC microbial community, has been detected by molecular methods in MFCs.32,34,35 However, there are fewer studies that describe CFB representatives. Here, we report for the first time successful isolation of a member of the phylum Bacteroidetes, strain TRS1-A1, similar to Marinifilum fragile, and we have demonstrated that this isolate can act as an exoelectrogenic bacterium. ε-Proteobacteria bacteria, such as Arcobacter species, were first isolated from an MFC in 2008,36 and the ability of members of this genus to produce electricity was demonstrated in 2009.13 In this study, strain MS30 was also confirmed as an electrogen, supporting these observations. The majority of Gram-negative bacteria can produce electricity.12,37 However, relatively few Gram-positive species are exoelectrogens.12 However, we report here power generation by B. stratosphericus (strain MS10), with sustained generation of electricity of 87.5 mW·m−2 in acetate-fed MFC. This is novel in the phylum Firmicutes. Furthermore, we have shown that members of the genera Vibrio, Enterobacter, and Citrobacter can act as exoelectrogens. Although Vibrio has been

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ASSOCIATED CONTENT

S Supporting Information *

One figure, showing a typical electrochemical impedance spectrum for a dual-chamber MFC. This material is available free of charge via the Internet at http://pubs.acs.org. 2990

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AUTHOR INFORMATION

Corresponding Author

*Phone: +44 (0)191 222 3057; e-mail: [email protected]. uk. Present Address ⊥

MRC Protein Phosphorylation Unit, College of Life Sciences, University of Dundee, Dundee, Scotland DD1 5EH, United Kingdom. Author Contributions ∥

J. Zhang and E. Zhang contributed equally to this work.

Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We thank Alexis Lau for his assistance with sediment sampling and for initially suggesting studies on bioelectricity due to his interest in electric fish, which led to this collaboration being formed. This work was also supported by a National Natural Science Foundation of China (NSFC) grant (20873120) to E.Z.. K.S. thanks EPSRC for financial support. J.G.B. thanks the BBSRC for the award of a Dorothy Hodgkin Studentship to J.Z. and the NERC for financial support.

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