- Email: [email protected]
Electron transfer pathways in microbial oxygen biocathodes
Electrochimica Acta 55 (2010) 813–818
Contents lists available at ScienceDirect
Electrochimica Acta journal homepage: www.elsevier.com/locate/electacta
Electron transfer pathways in microbial oxygen biocathodes Stefano Freguia ∗ , Seiya Tsujimura, Kenji Kano Bio-analytical and Physical Chemistry Laboratory, Division of Applied Life Sciences, Graduate School of Agriculture, Kyoto University, Sakyo-ku, Kyoto 606-8205, Japan
a r t i c l e
i n f o
Article history: Received 30 June 2009 Received in revised form 7 September 2009 Accepted 7 September 2009 Available online 15 September 2009 Keywords: Biocathode Extracellular electron transfer Insoluble electron donor Microbial fuel cell Oxygen reduction
a b s t r a c t The ability of some bacteria to enhance the rate of cathodic oxygen reduction to water has been recently discovered, opening the way to an entirely renewable and environmentally friendly concept of biocathode. In this study we reveal that several mechanisms may induce catalytic effects by bacteria. These comprise mechanisms that are putatively beneficial to the bacteria as well as mechanisms which are merely side effects, including quinone autoxidation and direct O2 reduction by heme compounds. Here we showed that 1 M of ACNQ is able to generate a significant catalytic wave for oxygen reduction, with onset at approximately 0 V vs. SHE. Similarly, adsorption of hemin on a carbon surface catalyses O2 reduction to H2 O2 with an onset of +0.2 V vs. SHE. To evaluate the catalytic pathways of live cells on cathodic oxygen reduction, two species of electrochemically active bacteria were selected as pure cultures, namely Acinetobacter calcoaceticus and Shewanella putrefaciens. The former appears to exploit a self-excreted redox compound with redox characteristics matching those of pyrroloquinoline quinone (PQQ) for extracellular electron transfer. The latter appears to utilise outer membrane-bound redox compounds. Interaction of quinones and cytochromes with the membrane-bound electron transfer chain is yet to be proven. © 2009 Elsevier Ltd. All rights reserved.
1. Introduction The problem of high overpotential for the cathodic reduction of oxygen to water is the main impediment to a simple and cheap implementation of chemical fuel cells as well as biofuel cells. While the thermodynamic potential for the reduction of O2 to water at pH 7 is 0.82 V vs. standard hydrogen electrode (SHE), the onset of reduction current on non-catalysed carbon is often below −0.1 V, depending on the type of carbon used. Catalysis is essential for this reaction and different strategies have been investigated and applied over the years. For chemical fuel cells (oxidizing energy-rich compounds such as methanol, methane or hydrogen), often precious metal catalysts such as platinum have been justified due to the high electricity throughput. However, reducing the installation costs by the use of cheaper catalysts is highly desirable. Moreover, often metal catalysis requires high temperatures to produce high current densities. Catalysts that work at ambient conditions would reduce the operating cost of the fuel cell. It was with this objective in mind that biofuel cells were devised, as processes that are able to couple the enzymatic oxidation of a variety of organic compounds to
∗ Corresponding author. Tel.: +81 75 753 6392; fax: +81 75 753 6456. E-mail addresses: [email protected] (S. Freguia), [email protected] (S. Tsujimura), [email protected] (K. Kano). 0013-4686/$ – see front matter © 2009 Elsevier Ltd. All rights reserved. doi:10.1016/j.electacta.2009.09.027
the enzymatic reduction of oxygen to water. Oxygen biocathodes originated with the discovery that fungal laccases exhibit catalytic effects on cathodic oxygen reduction, in the absence [1,2] and presence of mediators [3]. Several biocathodes have been developed in the past several years, all relying on the catalytic properties of multi-copper oxidases, including copper efflux oxidase (CueO) [4], bilirubin oxidase (BOD) [5] and laccases [6]. The enzymes work at ambient temperatures and often perform better than Pt as catalysts, thus becoming very attractive for this application. In particular, they can greatly reduce the overpotential of the oxygen reduction reaction, with onsets of catalytic current in the range of +0.6 to 0.7 V vs. SHE. Unfortunately, the complex proteic structure of these enzymes does not endure ambient conditions for very long, and denaturation imposes frequent catalyst replacements, with consequent considerable associated costs [7]. With the recent advent of microbial fuel cell research, a new concept of catalysis has been unveiled. Bacteria were first shown to catalyse the anodic oxidation of a wide array of organic compounds via the interaction of the electron transfer chain with the anode via a number of strategies, including excretion of soluble redox mediators [8], expression of outer membrane-bound cytochromes [9–11] and conductive pili, also known as nanowires [12]. A number of studies suggested that mixed cultures of bacteria growing at the cathode of microbial fuel cells are able to significantly reduce the overpotential of the four-electron reduction of O2 to water [13–16].
814
S. Freguia et al. / Electrochimica Acta 55 (2010) 813–818
The link between bacteria and catalysis was first shown by Rabaey et al. [17], who isolated pure cultures of bacteria that were shown to be active as cathode catalysts when allowed to grow on a cathode as a biofilm. Similarly to enzymes, bacteria can reduce the overpotential of the O2 reduction reaction, with catalytic current onsets as high as +0.45 V SHE [17,18]. As bacteria thrive at ambient conditions and neutral pH and can self-regenerate if properly supplied with a carbon source and nutrients, bacteria as catalysts have great potential to improve the performance and reduce the costs of biofuel cells. In order to improve and optimize microbial biocathodes, it is necessary to improve the understanding of their catalytic mechanisms. However, up to now no reports are available on the electron transfer mechanisms in microbial biocathodes. It was the objective of this work to shed some light on the possible electron transfer pathways exploited (or simply enabled) by bacteria at oxygen cathodes. For this purpose, we selected two strains of bacteria that were previously isolated in microbial fuel cells. Acinetobacter calcoaceticus was selected based on its prevalence in a mixed culture microbial fuel cell biocathode [17]. This bacterium is known for its ability to produce large amounts of pirroloquinoline quinone (PQQ), used by this bacterium as a prosthetic group for glucose dehydrogenase, a protein that exists both in membrane-spanning form and periplasmic soluble form [19]. Shewanella putrefaciens was chosen due its versatility in extracellular electron transfer mechanisms, in particular its known ability to locate large numbers and variety of cytochromes on the external side of the outer membrane [20]. S. putrefaciens is known to be able to perform anodic extracellular electron transfer via excreted riboflavin [21], outer membrane cytochromes [22] as well as nanowires [23]. In this work we used electrochemical techniques to analyze the nature of the electron transfer catalysis by these bacteria. However, unlike anodic organics oxidation, the reduction of oxygen to water (often via H2 O2 ) often can occur via an array of electrochemical reactions which may not involve the bacteria directly, but may exploit compounds produced by the bacteria. In particular, bacteria are known to produce and sometimes excrete quinones and heme-containing groups. It is known that quinone autoxidation is responsible for oxygen reduction to H2 O2 [24]. Hemin as well as catalase were also previously shown to catalyse cathodic oxygen reduction when adsorbed onto glassy carbon cathodes [25,26]. As the occurrence of these reactions may interfere with other truly bacterial catalytic pathways, we also included in this work the investigation of quinone and heme effects on the cathodic reduction of O2 . By investigating all the above mechanisms with the same electrochemical set-up and at the same conditions, a deeper understanding of the relative importance of these mechanisms at a biocathode becomes possible. 2. Experimental 2.1. Bacteria and culture conditions A. calcoaceticus (NBRC 12552) and S. putrefaciens (NBRC 3908) were obtained from the National Institute of Technology and Evaluation Biological Resource Center (Japan). Cultivation was carried out aerobically in 0.5-L flasks containing 250 mL of growth medium at 30 ◦ C on a shaker. A. calcoaceticus medium contained (per litre) polypeptone (10 g), yeast extract (2 g), MgSO4 ·7H2 O (1 g). S. putrefaciens medium contained (per litre) polypeptone (10 g), yeast extract (1 g), MgSO4 ·7H2 O (0.5 g) and Daigo artificial seawater (36 g). Cultures were harvested at late exponential phase (approximately 16 h after inoculation) via centrifugation at 5000 × g at 4 ◦ C, followed by 2 times wash in phosphate buffer saline (1.4 g L−1 KH2 PO4 , 7.6 g L−1 NaCl, pH 7.2) to eliminate all nutrients and metabolites from the cell wall.
2.2. Chemicals 2-Amino-3-dicarboxy-1,4-naphthoquinone (ACNQ) was obtained from Meiji Dairy Company (Japan). Hemin, PQQ and catalase were obtained from Sigma and used as received. 2.3. Electrochemical bioreactor The electrochemical bioreactor (previously detailed in Freguia et al. [27]) consisted of a cylindrical cell (14.5 cm height × 4.5 cm diameter) housing the working electrode (cathode). The counter electrode was placed in a second (smaller) cylinder, separated from the main chamber by a cellophane membrane (P-5 # 300, Futamura Chemical, Japan). The previously detailed [27] buffered nutrient medium (0.1 M phosphate buffer and nutrients, pH 7.0) was used as catholyte. The anolyte consisted of the same medium without nutrients. In this case the cathode was a 3 cm × 3 cm square of carbon paper (TGP-H-120, Toray, Japan). The choice of a small surface area cathode was dictated by the need to minimize background (electrochemical) oxygen reduction during the experiments. Current collection was guaranteed by a Ti mesh and wire. The reference electrode was in all cases a saturated Ag|AgCl electrode (−0.198 V vs SHE), inserted into the cathode compartment. Aeration was provided via filtered air supplied by an aquarium pump. Carbon source was not required for S. putrefaciens, whereas for A. calcoaceticus the biocathode cell was initially supplemented with 1.5 mM acetate to enable bacterial synthesis of redox active compounds. The cathode liquid volume was 0.1 L. The experiments were carried out in an incubator at the controlled temperature of 30 ± 1 ◦ C. Only for the tests of the electrochemical activity of bacterial cells harvested from the electrochemical bioreactor and their filtered supernatant, was carbon paper replaced with more reactive graphite felt (Osaka Gas) in a smaller 3-electrode one-compartment cell. 2.4. Chronoamperometry and cyclic voltammetry Following injection with pure cultures, the cathode (working electrode), anode (counter electrode) and reference electrode were connected to a potentiostat (HA-151A, Hokuto Denko, Tokyo, Japan). The potential of the anode was maintained at −200 mV vs. SHE. The current was continuously measured and recorded using a data acquisition unit (GL-800, GRAPHTEC, Yokohama, Japan). After full development of a catalytic current, cyclic voltammetry (CV) was performed directly on the setup using a potentiostat (ALS/CH Instruments, Austin, TX, USA). The cathode potential was swept from +400 to −300 mV SHE (unless otherwise stated) at scan rates from 10 to 40 mV s−1 . Cyclic voltammetry was done after stopping the chronoamperometric measurements, both in anaerobic conditions under Ar sparging (to reveal redox peaks) and under aeration (to reveal catalytic current). The experiment was repeated several times from different colonies of the bacteria to determine the reproducibility of the results. In each case, multiple scans were performed until the shape of the CVs was not changing appreciably (normally 2–4 times). For the tests with ACNQ, hemin and catalase, cyclic voltammetry was performed immediately after injection of the respective compound under aerated conditions (unless otherwise stated) and repeated several times over a period of 3 h to ensure that the observed effect was not due to temporary discharge. 3. Results 3.1. Effect on O2 reduction of ACNQ, hemin and catalase The addition of 1 M ACNQ to an abiotic cathode was followed by cyclic voltammetric analysis. A catalytic current was
S. Freguia et al. / Electrochimica Acta 55 (2010) 813–818
Fig. 1. Cyclic voltammograms of an abiotic carbon paper (9 cm2 ) cathode (in 0.1 M phosphate buffer, pH 7.0) in the presence of different compounds. Line A: catalase (0.1 mg L−1 ), aerated; line B: control (only phosphate buffer), aerated; line C: hemin (1 M) with 10 mM H2 O2 , anaerobic; line D: ACNQ (1 M), aerated; line E: hemin (1 M), aerated. Scan rate: 10 mV s−1 .
observed with onset corresponding to the reduction potential of ACNQ (−71 mV SHE), as shown by the cyclic voltammogram in Fig. 1 (line D). The significant difference with the ACNQ-free control (line B) can be attributed to quinone autoxidation. The inflection in the cyclic voltammogram at about −0.15 V SHE is typical of quinone autoxidations which proceed via semiquinone oxidation [24]. Quinone (zero-valent) and hydroquinone (bivalent) spontaneously generate some semiquinone (monovalent), whose oxidation by O2 is a one-electron process which generates superoxide radical O2 •− , which subsequently spontaneously disproportionate to O2 and H2 O2 . The net reaction is thus the twoelectron reduction of O2 to H2 O2 . More details on the quinone autoxidation process can be found in Tatsumi et al. [24]. In Fig. 1 the effect of hemin addition (1 M) is also shown (line E). It can be inferred that the heme group adsorbed onto the surface of carbon paper is able to bind and activate the oxygen molecule causing its reduction to H2 O2 with onset at around +0.1 V SHE. The reduction mostly stops at hydrogen peroxide, as only minor catalytic effect was observed upon injecting 10 mM H2 O2 into an anaerobic hemin cathode (line C). Despite previous reports that catalase directly enables O2 reduction to water at glassy carbon surfaces [26], here the addition of 0.1 mg L−1 catalase on carbon paper not only did not exhibit catalytic effects, but also slowed down the non-catalysed O2 reduction on the carbon surface, possibly by reducing the available surface area due to catalase adsorption (Fig. 1, line A).
815
Fig. 2. Chronoamperometry (−200 mV vs. SHE) of carbon paper aerated biocathodes (9 cm2 , initial pH 7.0, continuously aerated). Solid line: A. calcoaceticus biocathode; dashed line: S. putrefaciens biocathode. The gaps in the line indicate interruptions of chronoamperometry to perform cyclic voltammetry.
Faradaic and non-Faradaic current dependences on the scan rate, [28]). As the mid-point potential corresponds well to that of PQQ (based on direct comparison with PQQ peak on carbon paper as shown in Fig. 4B) and based on well established knowledge that this bacterium produces significant amounts of this cofactor [19], tests were carried out to establish the ability of A. calcoaceticus to exploit exogenous PQQ for extracellular electron uptake from the cathode. The addition of 2 M PQQ produced a steeper catalytic current in cyclic voltammetry (Fig. 3, line D), thus indicating that the bacterium exploited the extra mediator for oxygen reduction. Multiple additions of PQQ up to 4.5 M were carried out during chronoamperometry at −200 mV SHE (Fig. 5), showing a proportional increase of the current with the exogenous PQQ concentration. Therefore it appears likely that PQQ is the redox compound produced and utilised by the bacterium for extracellular electron transfer. A second larger reversible peak was detected at much lower potential (around −0.27 V vs. SHE, see Fig. 4B) and was likely responsible for the increased catalytic O2 reduction at low potential (<0.2 V vs. SHE). Peak height analysis as shown in Fig. 4B (lower inset) suggested that this redox compound may be adsorbed onto the cathode surface, in light of the linear dependence of the peak height on the scan rate. As the compound should be linked to the bacterial membrane as well in order to be catalytically active, it is possible that it consists of a membrane-bound protein directly interacting with the electrode. Despite being interesting on the
3.2. A. calcoaceticus catalyses O2 reduction via electron shuttling by a diffusive mediator After approximately 3 h following the injection of the A. calcoaceticus harvested cells into the electrochemical bioreactor, a catalytic current developed as indicated by chronoamperometry (Fig. 2). Cyclic voltammetry revealed that the onset of the catalytic effect occurred at slightly less than +0.1 V vs. SHE (Fig. 3, line C). Cyclic voltammograms obtained under anaerobic conditions (Fig. 4A) revealed that a reversible peak was present with a mid-point potential of approximately +60 mV vs. SHE and this peak corresponded to the onset of catalytic current. Peak height analysis (Fig. 4B, upper inset) showed that the peak height was proportional to the square root of the scan rate, suggesting that a diffusive mediator was responsible for this catalysis (however, this technique may produce misleading results as a consequence of the difficulty of selecting a background current for the peaks, as well as due to the influence of electrode porosity on both the
Fig. 3. Cyclic voltammograms of a carbon paper cathode (9 cm2 ) in aerated buffered medium (pH 7.0). Line A: control (only phosphate buffer); line B: PQQ (2 M); line C: A. calcoaceticus; line D: A. calcoaceticus and 2 M PQQ. Scan rate: 10 mV s−1 .
816
S. Freguia et al. / Electrochimica Acta 55 (2010) 813–818
Fig. 6. Cyclic voltammograms of a carbon paper biocathode (9 cm2 , pH 7.0, aerated). Line A: abiotic control; line B: S. putrefaciens biocathode. Inset: cyclic voltammograms of a graphite felt biocathode (pH 7.0, air saturated). Dotted line: abiotic control; solid line: double washed S. putrefaciens cells harvested from a carbon paper biocathode; dashed line: filtered supernatant from the same carbon paper biocathode. Scan rate: 10 mV s−1 .
Fig. 4. (A) Cyclic voltammograms displaying the onset of catalytic current for an A. calcoaceticus carbon paper biocathode (9 cm2 , pH 7.0). Solid line: anaerobic conditions, displaying a reversible peak; dashed line: aerated conditions, showing that the onset of catalytic current corresponds to the potential of the redox peak. Scan rate: 10 mV s−1 . (B) Peak height analysis of an A. calcoaceticus biocathode (carbon paper, pH 7.0, anaerobic conditions). Solid lines, from inner to outer: cyclic voltammograms at scan rates of 10, 20 and 40 mV s−1 . Dashed line: addition of 2 mM PQQ, scan rate of 10 mV s−1 . Insets: peak height vs. square root of scan rate for the high potential peak (putative PQQ) and peak height vs. scan rate for the low potential peak.
electron transfer mechanism point of view, the redox potential of this compound is too low to make it a useful cathodic catalyst in microbial fuel cells (as the anode potential in microbial fuel cells is normally higher than −0.27 V vs. SHE). To evaluate the possibility that the observed catalytic current might be due only to quinone autoxidation, cyclic voltammetry was carried out with a control electrode supplied only with buffer and 2 M PQQ (Fig. 3, line B). The minor effect of PQQ on the abiotic electrode indicated that, unlike ACNQ, PQQ is not prone to autoxidation and that the increased oxygen reduction current must be attributed directly to the bacterium. This small rate of autoxidation for PQQ matches well the values measured by Itoh et al. [29], in the absence of significant concentrations of Ca2+ . Moreover, cyclic voltammetry of the filtered A. calcoaceticus catholyte did not reveal any catalytic effect (data not shown), indicating the absence of other extracellular abiotic catalytic processes at this biocathode. 3.3. S. putrefaciens catalyses O2 reduction via membrane associated compounds
Fig. 5. Chronoamperometry (−200 mV vs. SHE) of a carbon paper A. calcoaceticus biocathode (9 cm2 , pH 7.0, aerated) supplied with different concentrations of exogenous PQQ: the arrows indicate PQQ injection points (1.5 M per injection). The increase in catalytic current is roughly proportional to the concentration of added PQQ.
The development of catalytic current of oxygen reduction by S. putrefaciens can be followed in Fig. 2. The longer time required for the development of catalytic current compared to A. calcoaceticus is attributed to large amounts of carbon storage by S. putrefaciens (as revealed by 72 h long anodic current production by washed Shewanella cell in anodic experiments, data not shown). After 24 h of chronoamperometry at −200 mV SHE, cyclic voltammetry revealed a catalytic wave with onset at slightly less than +0.2 V SHE (Fig. 6). In the case of Shewanella, no peaks could be observed under anaerobic conditions, suggesting that possibly the last step of electron transfer is not carried out by a reversible redox compound. To determine the nature of the electron transfer, Shewanella cells were harvested from the bioelectrochemical reactor, washed twice with phosphate buffer saline, resuspended in phosphate buffer and immediately supplied to a pyrolytic graphite felt electrode with high surface area and activity for quick determination of the electrochemical response. In air-saturated conditions, a catalytic current with similar onset was observed (inset of Fig. 6), suggesting that redox compounds located on the external side of the outer membrane may be responsible for the last step of electron transfer in this case. The onset of the catalytic current is indeed close to the potential of some of the outer membrane cytochromes previously extracted from S. putrefaciens, including OmcA and
S. Freguia et al. / Electrochimica Acta 55 (2010) 813–818
Fig. 7. Close up of the onset of catalytic current as observed by cyclic voltammetry of carbon paper cathodes (9 cm2 ) at pH 7.0 and aerated conditions. Dotted line: abiotic control; solid line: S. putrefaciens biocathode; dashed line: abiotic cathode supplemented with 1 M hemin. Scan rate: 10 mV s−1 .
MtrC [30]. However, the filtered supernatant of the harvested cells exhibited exactly the same cyclic voltammetry shape and magnitude (see inset of Fig. 6), indicating that the compounds present on the surface of Shewanella’s outer membrane may also be released into the surrounding medium, where they may be able to carry out an abiotic catalytic effect similar to that of free hemin. A comparison of the onsets of the catalytic currents by Shewanella cells and hemin is shown in Fig. 7, revealing identical onsets for the two cases. The catalytic effect by the filtered supernatant may also be due to hemes released due to cell lysis. 4. Discussion Despite the possibility that oxygen reduction might occur based on non-biological pathways such as the ones described above (quinone autoxidation and direct electrochemistry of hemes), pathways that do require live bacteria are also present. The results obtained for A. calcoaceticus suggest the involvement of free PQQ in the catalysis. While PQQ autoxidation is not happening at significant rates, when the cofactor is supplied to the Acinetobacter biocathode, it is able to enhance the rate of cathodic oxygen reduction. Thus, there must be a pathway that involves the bacterium directly as a catalyst. Even though PQQ is normally bound to glucose dehydrogenase as a cofactor in Acinetobacter [19], pathways in which PQQ shuttles electrons between the soluble enzyme and an electrode have been previously proposed [31]. As the natural electron acceptor of PQQ is cytochrome c [19], it is likely that once PQQ is reduced by the cathode, electrons are transferred to cytochrome c. Finally cytochrome oxidases may complete the electron transfer to dioxygen, which is reduced to water. If we accept that PQQ is shuttling electrons between a cathode and the membranebound electron transfer chain, this proposed pathway becomes an easy possibility, which moreover would provide the bacteria with the important advantage that due to proton translocation during cytochrome c oxidation by complex IV, ATP may be produced through this reaction. In this case, bacteria would effectively use a cathode as insoluble electron donor for energy generation. While the results of this preliminary work suggest that this may be the case, proving that this indeed happens would require testing with cytochrome c and PQQ-dependent glucose dehydrogenase deletion mutants. However, in support of this theory, recent findings that bacteria can reduce nitrate (NO3 − ) to nitrite or N2 using a cathode as electron donor also point in this direction [32,33]. As nitrate reduction is not subject to enzyme-free reactions like O2 , it is most likely that in the case of nitrate reduction bacteria are exploiting
817
the electron transfer chain to deliver electrons from a cathode to the terminal electron acceptor. In the case of S. putrefaciens, membrane attached compounds appear to generate the catalytic current. As the bacterium is well known to localize c-type cytochromes on the external surface of the outer membrane, it is likely that cytochromes or cytochromecontaining nanowires are interacting directly with the cathode. However, in this case the electron transfer chain may be partially by-passed as oxygen may spontaneously react with hemes on the outer surface of the bacterium. The onset of oxygen reduction current with the bacteria tested in this study is at most +0.1 V vs. SHE. In two independent mixed culture studies [17,18], onsets as high as +0.45 V were obtained. More work needs to be done to isolate the bacteria responsible for such high onset of catalytic current, as well as to identify the bacterial redox compounds that enable such extracellular electron transfer. While enzymatic biocathodes can certainly achieve lower overpotentials with onsets of catalytic current as high as +0.75 V vs. SHE, the advantages of a self-renewable bacterial catalyst are numerous, justifying further research in this direction. 5. Conclusions The catalysis of oxygen reduction at microbial biocathodes may occur via several parallel pathways. Some of these pathways may be linked to the metabolic activity of the bacteria, while others are a side effect of bacterial excretion of redox active compounds which have the ability to chemically react with oxygen. Among redox active metabolites which are normally produced and excreted by bacteria, quinones in particular undergo the phenomenon of autoxidation, reducing O2 to O2 •− in solution and subsequently getting reduced at the cathode surface. Free hemes which may be excreted directly or exist in solution as lysis products have instead the ability to directly reduce O2 to H2 O2 at the cathode surface. The extracellular electron pathways which require the metabolic activity of the bacteria directly are likely to involve the electrical interaction between the cathode and the bacteria’s electron transfer chain. This interaction may be carried out via shuttling of excreted quinones, or via direct contact by redox active compounds (such as c-type cytochromes) located on the outer membrane of the bacteria. Acknowledgement The first author was supported by the Japan Society for the Promotion of Science (PE 08026). References [1] C.W. Lee, H.B. Gray, F.C. Anson, B.G. Malmstrom, J. Electroanal. Chem. (1984) 289. [2] M.R. Tarasevich, A.I. Yaropolov, V.A. Bogdanovskaya, S.D. Varfolomeev, Bioelectrochem. Bioenerg. 6 (1979) 393. [3] G.T.R. Kim, H.H. Palmore, J. Electroanal. Chem. 464 (1999) 110. [4] S. Tsujimura, Y. Miura, K. Kano, Electrochim. Acta 53 (2008) 5716. [5] S. Tsujimura, B. Tatsumi, J. Ogawa, S. Shimizu, K. Kano, T. Ikeda, J. Electroanal. Chem. 496 (2001) 69. [6] S. Tsujimura, Y. Kamitaka, K. Kano, Fuel Cells 7 (2007) 463. [7] C. Vaz-Dominguez, S. Campuzano, O. Rudiger, M. Pita, M. Gorbacheva, S. Shleev, V.M. Fernandez, A.L. De Lacey, Biosens. Bioelectron. 24 (2008) 531. [8] K. Rabaey, N. Boon, S.D. Siciliano, M. Verhaege, W. Verstraete, Appl. Environ. Microbiol. 70 (2004) 5373. [9] D.E. Holmes, S.K. Chaudhuri, K.P. Nevin, T. Mehta, B.A. Methe, A. Liu, J.E. Ward, T.L. Woodard, J. Webster, D.R. Lovley, Environ. Microbiol. 8 (2006) 1805. [10] B.H. Kim, T. Ikeda, H.S. Park, H.J. Kim, M.S. Hyun, K. Kano, K. Takagi, H. Tatsumi, Biotechnol. Tech. 13 (1999) 475. [11] H. Richter, K.P. Nevin, H.F. Jia, D.A. Lowy, D.R. Lovley, L.M. Tender, Energy Environ. Sci. 2 (2009) 506.
818
S. Freguia et al. / Electrochimica Acta 55 (2010) 813–818
[12] G. Reguera, K.D. McCarthy, T. Mehta, J.S. Nicoll, M.T. Tuominen, D.R. Lovley, Nature 435 (2005) 1098. [13] A. Bergel, D. Feron, A. Mollica, Electrochem. Commun. 7 (2005) 900. [14] P. Clauwaert, D. Van der Ha, N. Boon, K. Verbeken, M. Verhaege, K. Rabaey, W. Verstraete, Environ. Sci. Technol. 41 (2007) 7564. [15] S. Freguia, K. Rabaey, Z. Yuan, J. Keller, Water Res. 42 (2008) 1387. [16] V. Scotto, R. Dicintio, G. Marcenaro, Corros. Sci. 25 (1985) 185. [17] K. Rabaey, S.T. Read, P. Clauwaert, S. Freguia, P.L. Bond, L.L. Blackall, J. Keller, ISME J. 2 (2008) 519. [18] A. Aldrovandi, E. Marsili, L. Stante, P. Paganin, S. Tabacchioni, A. Giordano, Bioresour. Technol. 100 (2009) 3252. [19] K. Matsushita, H. Toyama, M. Yamada, O. Adachi, Appl. Microbiol. Biotechnol. 58 (2002) 13. [20] C.R. Myers, J.M. Myers, J. Bacteriol. 174 (1992) 3429. [21] E. Marsili, D.B. Baron, I.D. Shikhare, D. Coursolle, J.A. Gralnick, D.R. Bond, Proc. Natl. Acad. Sci. U.S.A. 105 (2008) 3968. [22] H.J. Kim, H.S. Park, M.S. Hyun, I.S. Chang, M. Kim, B.H. Kim, Enzyme Microb. Technol. 30 (2002) 145.
[23] Y.A. Gorby, S. Yanina, J.S. McLean, K.M. Rosso, D. Moyles, A. Dohnalkova, T.J. Beveridge, I.S. Chang, B.H. Kim, K.S. Kim, D.E. Culley, S.B. Reed, M.F. Romine, D.A. Saffarini, E.A. Hill, L. Shi, D.A. Elias, D.W. Kennedy, G. Pinchuk, K. Watanabe, S. Ishii, B. Logan, K.H. Nealson, J.K. Fredrickson, Proc. Natl. Acad. Sci. U.S.A. 103 (2006) 11358. [24] H. Tatsumi, H. Nakase, K. Kano, T. Ikeda, J. Electroanal. Chem. 443 (1998) 236. [25] H. Iken, L. Etcheverry, A. Bergel, R. Basseguy, Electrochim. Acta 54 (2008) 60. [26] Bergel v, A. Lai, J. Electroanal. Chem. 494 (2000) 30. [27] S. Freguia, M. Masuda, S. Tsujimura, K. Kano, Bioelectrochemistry 76 (2009) 14. [28] D. Compton, R.G. Menshykau, Electroanalysis 20 (2008) 2387. [29] S. Itoh, H. Kawakami, S. Fukuzumi, Chem. Commun. (1997) 29. [30] M. Firer-Sherwood, G.S. Pulcu, S.J. Elliott, J. Biol. Inorg. Chem. 13 (2008) 849. [31] V. Laurinavicius, J. Razumiene, A. Ramanavicius, A.D. Ryabov, Biosens. Bioelectron. 20 (2004) 1217. [32] K.B. Gregory, D.R. Bond, D.R. Lovley, Environ. Microbiol. 6 (2004) 596. [33] B. Virdis, K. Rabaey, Z. Yuan, J. Keller, Water Res. 42 (2008) 3013.
Contents lists available at ScienceDirect
Electrochimica Acta journal homepage: www.elsevier.com/locate/electacta
Electron transfer pathways in microbial oxygen biocathodes Stefano Freguia ∗ , Seiya Tsujimura, Kenji Kano Bio-analytical and Physical Chemistry Laboratory, Division of Applied Life Sciences, Graduate School of Agriculture, Kyoto University, Sakyo-ku, Kyoto 606-8205, Japan
a r t i c l e
i n f o
Article history: Received 30 June 2009 Received in revised form 7 September 2009 Accepted 7 September 2009 Available online 15 September 2009 Keywords: Biocathode Extracellular electron transfer Insoluble electron donor Microbial fuel cell Oxygen reduction
a b s t r a c t The ability of some bacteria to enhance the rate of cathodic oxygen reduction to water has been recently discovered, opening the way to an entirely renewable and environmentally friendly concept of biocathode. In this study we reveal that several mechanisms may induce catalytic effects by bacteria. These comprise mechanisms that are putatively beneficial to the bacteria as well as mechanisms which are merely side effects, including quinone autoxidation and direct O2 reduction by heme compounds. Here we showed that 1 M of ACNQ is able to generate a significant catalytic wave for oxygen reduction, with onset at approximately 0 V vs. SHE. Similarly, adsorption of hemin on a carbon surface catalyses O2 reduction to H2 O2 with an onset of +0.2 V vs. SHE. To evaluate the catalytic pathways of live cells on cathodic oxygen reduction, two species of electrochemically active bacteria were selected as pure cultures, namely Acinetobacter calcoaceticus and Shewanella putrefaciens. The former appears to exploit a self-excreted redox compound with redox characteristics matching those of pyrroloquinoline quinone (PQQ) for extracellular electron transfer. The latter appears to utilise outer membrane-bound redox compounds. Interaction of quinones and cytochromes with the membrane-bound electron transfer chain is yet to be proven. © 2009 Elsevier Ltd. All rights reserved.
1. Introduction The problem of high overpotential for the cathodic reduction of oxygen to water is the main impediment to a simple and cheap implementation of chemical fuel cells as well as biofuel cells. While the thermodynamic potential for the reduction of O2 to water at pH 7 is 0.82 V vs. standard hydrogen electrode (SHE), the onset of reduction current on non-catalysed carbon is often below −0.1 V, depending on the type of carbon used. Catalysis is essential for this reaction and different strategies have been investigated and applied over the years. For chemical fuel cells (oxidizing energy-rich compounds such as methanol, methane or hydrogen), often precious metal catalysts such as platinum have been justified due to the high electricity throughput. However, reducing the installation costs by the use of cheaper catalysts is highly desirable. Moreover, often metal catalysis requires high temperatures to produce high current densities. Catalysts that work at ambient conditions would reduce the operating cost of the fuel cell. It was with this objective in mind that biofuel cells were devised, as processes that are able to couple the enzymatic oxidation of a variety of organic compounds to
∗ Corresponding author. Tel.: +81 75 753 6392; fax: +81 75 753 6456. E-mail addresses: [email protected] (S. Freguia), [email protected] (S. Tsujimura), [email protected] (K. Kano). 0013-4686/$ – see front matter © 2009 Elsevier Ltd. All rights reserved. doi:10.1016/j.electacta.2009.09.027
the enzymatic reduction of oxygen to water. Oxygen biocathodes originated with the discovery that fungal laccases exhibit catalytic effects on cathodic oxygen reduction, in the absence [1,2] and presence of mediators [3]. Several biocathodes have been developed in the past several years, all relying on the catalytic properties of multi-copper oxidases, including copper efflux oxidase (CueO) [4], bilirubin oxidase (BOD) [5] and laccases [6]. The enzymes work at ambient temperatures and often perform better than Pt as catalysts, thus becoming very attractive for this application. In particular, they can greatly reduce the overpotential of the oxygen reduction reaction, with onsets of catalytic current in the range of +0.6 to 0.7 V vs. SHE. Unfortunately, the complex proteic structure of these enzymes does not endure ambient conditions for very long, and denaturation imposes frequent catalyst replacements, with consequent considerable associated costs [7]. With the recent advent of microbial fuel cell research, a new concept of catalysis has been unveiled. Bacteria were first shown to catalyse the anodic oxidation of a wide array of organic compounds via the interaction of the electron transfer chain with the anode via a number of strategies, including excretion of soluble redox mediators [8], expression of outer membrane-bound cytochromes [9–11] and conductive pili, also known as nanowires [12]. A number of studies suggested that mixed cultures of bacteria growing at the cathode of microbial fuel cells are able to significantly reduce the overpotential of the four-electron reduction of O2 to water [13–16].
814
S. Freguia et al. / Electrochimica Acta 55 (2010) 813–818
The link between bacteria and catalysis was first shown by Rabaey et al. [17], who isolated pure cultures of bacteria that were shown to be active as cathode catalysts when allowed to grow on a cathode as a biofilm. Similarly to enzymes, bacteria can reduce the overpotential of the O2 reduction reaction, with catalytic current onsets as high as +0.45 V SHE [17,18]. As bacteria thrive at ambient conditions and neutral pH and can self-regenerate if properly supplied with a carbon source and nutrients, bacteria as catalysts have great potential to improve the performance and reduce the costs of biofuel cells. In order to improve and optimize microbial biocathodes, it is necessary to improve the understanding of their catalytic mechanisms. However, up to now no reports are available on the electron transfer mechanisms in microbial biocathodes. It was the objective of this work to shed some light on the possible electron transfer pathways exploited (or simply enabled) by bacteria at oxygen cathodes. For this purpose, we selected two strains of bacteria that were previously isolated in microbial fuel cells. Acinetobacter calcoaceticus was selected based on its prevalence in a mixed culture microbial fuel cell biocathode [17]. This bacterium is known for its ability to produce large amounts of pirroloquinoline quinone (PQQ), used by this bacterium as a prosthetic group for glucose dehydrogenase, a protein that exists both in membrane-spanning form and periplasmic soluble form [19]. Shewanella putrefaciens was chosen due its versatility in extracellular electron transfer mechanisms, in particular its known ability to locate large numbers and variety of cytochromes on the external side of the outer membrane [20]. S. putrefaciens is known to be able to perform anodic extracellular electron transfer via excreted riboflavin [21], outer membrane cytochromes [22] as well as nanowires [23]. In this work we used electrochemical techniques to analyze the nature of the electron transfer catalysis by these bacteria. However, unlike anodic organics oxidation, the reduction of oxygen to water (often via H2 O2 ) often can occur via an array of electrochemical reactions which may not involve the bacteria directly, but may exploit compounds produced by the bacteria. In particular, bacteria are known to produce and sometimes excrete quinones and heme-containing groups. It is known that quinone autoxidation is responsible for oxygen reduction to H2 O2 [24]. Hemin as well as catalase were also previously shown to catalyse cathodic oxygen reduction when adsorbed onto glassy carbon cathodes [25,26]. As the occurrence of these reactions may interfere with other truly bacterial catalytic pathways, we also included in this work the investigation of quinone and heme effects on the cathodic reduction of O2 . By investigating all the above mechanisms with the same electrochemical set-up and at the same conditions, a deeper understanding of the relative importance of these mechanisms at a biocathode becomes possible. 2. Experimental 2.1. Bacteria and culture conditions A. calcoaceticus (NBRC 12552) and S. putrefaciens (NBRC 3908) were obtained from the National Institute of Technology and Evaluation Biological Resource Center (Japan). Cultivation was carried out aerobically in 0.5-L flasks containing 250 mL of growth medium at 30 ◦ C on a shaker. A. calcoaceticus medium contained (per litre) polypeptone (10 g), yeast extract (2 g), MgSO4 ·7H2 O (1 g). S. putrefaciens medium contained (per litre) polypeptone (10 g), yeast extract (1 g), MgSO4 ·7H2 O (0.5 g) and Daigo artificial seawater (36 g). Cultures were harvested at late exponential phase (approximately 16 h after inoculation) via centrifugation at 5000 × g at 4 ◦ C, followed by 2 times wash in phosphate buffer saline (1.4 g L−1 KH2 PO4 , 7.6 g L−1 NaCl, pH 7.2) to eliminate all nutrients and metabolites from the cell wall.
2.2. Chemicals 2-Amino-3-dicarboxy-1,4-naphthoquinone (ACNQ) was obtained from Meiji Dairy Company (Japan). Hemin, PQQ and catalase were obtained from Sigma and used as received. 2.3. Electrochemical bioreactor The electrochemical bioreactor (previously detailed in Freguia et al. [27]) consisted of a cylindrical cell (14.5 cm height × 4.5 cm diameter) housing the working electrode (cathode). The counter electrode was placed in a second (smaller) cylinder, separated from the main chamber by a cellophane membrane (P-5 # 300, Futamura Chemical, Japan). The previously detailed [27] buffered nutrient medium (0.1 M phosphate buffer and nutrients, pH 7.0) was used as catholyte. The anolyte consisted of the same medium without nutrients. In this case the cathode was a 3 cm × 3 cm square of carbon paper (TGP-H-120, Toray, Japan). The choice of a small surface area cathode was dictated by the need to minimize background (electrochemical) oxygen reduction during the experiments. Current collection was guaranteed by a Ti mesh and wire. The reference electrode was in all cases a saturated Ag|AgCl electrode (−0.198 V vs SHE), inserted into the cathode compartment. Aeration was provided via filtered air supplied by an aquarium pump. Carbon source was not required for S. putrefaciens, whereas for A. calcoaceticus the biocathode cell was initially supplemented with 1.5 mM acetate to enable bacterial synthesis of redox active compounds. The cathode liquid volume was 0.1 L. The experiments were carried out in an incubator at the controlled temperature of 30 ± 1 ◦ C. Only for the tests of the electrochemical activity of bacterial cells harvested from the electrochemical bioreactor and their filtered supernatant, was carbon paper replaced with more reactive graphite felt (Osaka Gas) in a smaller 3-electrode one-compartment cell. 2.4. Chronoamperometry and cyclic voltammetry Following injection with pure cultures, the cathode (working electrode), anode (counter electrode) and reference electrode were connected to a potentiostat (HA-151A, Hokuto Denko, Tokyo, Japan). The potential of the anode was maintained at −200 mV vs. SHE. The current was continuously measured and recorded using a data acquisition unit (GL-800, GRAPHTEC, Yokohama, Japan). After full development of a catalytic current, cyclic voltammetry (CV) was performed directly on the setup using a potentiostat (ALS/CH Instruments, Austin, TX, USA). The cathode potential was swept from +400 to −300 mV SHE (unless otherwise stated) at scan rates from 10 to 40 mV s−1 . Cyclic voltammetry was done after stopping the chronoamperometric measurements, both in anaerobic conditions under Ar sparging (to reveal redox peaks) and under aeration (to reveal catalytic current). The experiment was repeated several times from different colonies of the bacteria to determine the reproducibility of the results. In each case, multiple scans were performed until the shape of the CVs was not changing appreciably (normally 2–4 times). For the tests with ACNQ, hemin and catalase, cyclic voltammetry was performed immediately after injection of the respective compound under aerated conditions (unless otherwise stated) and repeated several times over a period of 3 h to ensure that the observed effect was not due to temporary discharge. 3. Results 3.1. Effect on O2 reduction of ACNQ, hemin and catalase The addition of 1 M ACNQ to an abiotic cathode was followed by cyclic voltammetric analysis. A catalytic current was
S. Freguia et al. / Electrochimica Acta 55 (2010) 813–818
Fig. 1. Cyclic voltammograms of an abiotic carbon paper (9 cm2 ) cathode (in 0.1 M phosphate buffer, pH 7.0) in the presence of different compounds. Line A: catalase (0.1 mg L−1 ), aerated; line B: control (only phosphate buffer), aerated; line C: hemin (1 M) with 10 mM H2 O2 , anaerobic; line D: ACNQ (1 M), aerated; line E: hemin (1 M), aerated. Scan rate: 10 mV s−1 .
observed with onset corresponding to the reduction potential of ACNQ (−71 mV SHE), as shown by the cyclic voltammogram in Fig. 1 (line D). The significant difference with the ACNQ-free control (line B) can be attributed to quinone autoxidation. The inflection in the cyclic voltammogram at about −0.15 V SHE is typical of quinone autoxidations which proceed via semiquinone oxidation [24]. Quinone (zero-valent) and hydroquinone (bivalent) spontaneously generate some semiquinone (monovalent), whose oxidation by O2 is a one-electron process which generates superoxide radical O2 •− , which subsequently spontaneously disproportionate to O2 and H2 O2 . The net reaction is thus the twoelectron reduction of O2 to H2 O2 . More details on the quinone autoxidation process can be found in Tatsumi et al. [24]. In Fig. 1 the effect of hemin addition (1 M) is also shown (line E). It can be inferred that the heme group adsorbed onto the surface of carbon paper is able to bind and activate the oxygen molecule causing its reduction to H2 O2 with onset at around +0.1 V SHE. The reduction mostly stops at hydrogen peroxide, as only minor catalytic effect was observed upon injecting 10 mM H2 O2 into an anaerobic hemin cathode (line C). Despite previous reports that catalase directly enables O2 reduction to water at glassy carbon surfaces [26], here the addition of 0.1 mg L−1 catalase on carbon paper not only did not exhibit catalytic effects, but also slowed down the non-catalysed O2 reduction on the carbon surface, possibly by reducing the available surface area due to catalase adsorption (Fig. 1, line A).
815
Fig. 2. Chronoamperometry (−200 mV vs. SHE) of carbon paper aerated biocathodes (9 cm2 , initial pH 7.0, continuously aerated). Solid line: A. calcoaceticus biocathode; dashed line: S. putrefaciens biocathode. The gaps in the line indicate interruptions of chronoamperometry to perform cyclic voltammetry.
Faradaic and non-Faradaic current dependences on the scan rate, [28]). As the mid-point potential corresponds well to that of PQQ (based on direct comparison with PQQ peak on carbon paper as shown in Fig. 4B) and based on well established knowledge that this bacterium produces significant amounts of this cofactor [19], tests were carried out to establish the ability of A. calcoaceticus to exploit exogenous PQQ for extracellular electron uptake from the cathode. The addition of 2 M PQQ produced a steeper catalytic current in cyclic voltammetry (Fig. 3, line D), thus indicating that the bacterium exploited the extra mediator for oxygen reduction. Multiple additions of PQQ up to 4.5 M were carried out during chronoamperometry at −200 mV SHE (Fig. 5), showing a proportional increase of the current with the exogenous PQQ concentration. Therefore it appears likely that PQQ is the redox compound produced and utilised by the bacterium for extracellular electron transfer. A second larger reversible peak was detected at much lower potential (around −0.27 V vs. SHE, see Fig. 4B) and was likely responsible for the increased catalytic O2 reduction at low potential (<0.2 V vs. SHE). Peak height analysis as shown in Fig. 4B (lower inset) suggested that this redox compound may be adsorbed onto the cathode surface, in light of the linear dependence of the peak height on the scan rate. As the compound should be linked to the bacterial membrane as well in order to be catalytically active, it is possible that it consists of a membrane-bound protein directly interacting with the electrode. Despite being interesting on the
3.2. A. calcoaceticus catalyses O2 reduction via electron shuttling by a diffusive mediator After approximately 3 h following the injection of the A. calcoaceticus harvested cells into the electrochemical bioreactor, a catalytic current developed as indicated by chronoamperometry (Fig. 2). Cyclic voltammetry revealed that the onset of the catalytic effect occurred at slightly less than +0.1 V vs. SHE (Fig. 3, line C). Cyclic voltammograms obtained under anaerobic conditions (Fig. 4A) revealed that a reversible peak was present with a mid-point potential of approximately +60 mV vs. SHE and this peak corresponded to the onset of catalytic current. Peak height analysis (Fig. 4B, upper inset) showed that the peak height was proportional to the square root of the scan rate, suggesting that a diffusive mediator was responsible for this catalysis (however, this technique may produce misleading results as a consequence of the difficulty of selecting a background current for the peaks, as well as due to the influence of electrode porosity on both the
Fig. 3. Cyclic voltammograms of a carbon paper cathode (9 cm2 ) in aerated buffered medium (pH 7.0). Line A: control (only phosphate buffer); line B: PQQ (2 M); line C: A. calcoaceticus; line D: A. calcoaceticus and 2 M PQQ. Scan rate: 10 mV s−1 .
816
S. Freguia et al. / Electrochimica Acta 55 (2010) 813–818
Fig. 6. Cyclic voltammograms of a carbon paper biocathode (9 cm2 , pH 7.0, aerated). Line A: abiotic control; line B: S. putrefaciens biocathode. Inset: cyclic voltammograms of a graphite felt biocathode (pH 7.0, air saturated). Dotted line: abiotic control; solid line: double washed S. putrefaciens cells harvested from a carbon paper biocathode; dashed line: filtered supernatant from the same carbon paper biocathode. Scan rate: 10 mV s−1 .
Fig. 4. (A) Cyclic voltammograms displaying the onset of catalytic current for an A. calcoaceticus carbon paper biocathode (9 cm2 , pH 7.0). Solid line: anaerobic conditions, displaying a reversible peak; dashed line: aerated conditions, showing that the onset of catalytic current corresponds to the potential of the redox peak. Scan rate: 10 mV s−1 . (B) Peak height analysis of an A. calcoaceticus biocathode (carbon paper, pH 7.0, anaerobic conditions). Solid lines, from inner to outer: cyclic voltammograms at scan rates of 10, 20 and 40 mV s−1 . Dashed line: addition of 2 mM PQQ, scan rate of 10 mV s−1 . Insets: peak height vs. square root of scan rate for the high potential peak (putative PQQ) and peak height vs. scan rate for the low potential peak.
electron transfer mechanism point of view, the redox potential of this compound is too low to make it a useful cathodic catalyst in microbial fuel cells (as the anode potential in microbial fuel cells is normally higher than −0.27 V vs. SHE). To evaluate the possibility that the observed catalytic current might be due only to quinone autoxidation, cyclic voltammetry was carried out with a control electrode supplied only with buffer and 2 M PQQ (Fig. 3, line B). The minor effect of PQQ on the abiotic electrode indicated that, unlike ACNQ, PQQ is not prone to autoxidation and that the increased oxygen reduction current must be attributed directly to the bacterium. This small rate of autoxidation for PQQ matches well the values measured by Itoh et al. [29], in the absence of significant concentrations of Ca2+ . Moreover, cyclic voltammetry of the filtered A. calcoaceticus catholyte did not reveal any catalytic effect (data not shown), indicating the absence of other extracellular abiotic catalytic processes at this biocathode. 3.3. S. putrefaciens catalyses O2 reduction via membrane associated compounds
Fig. 5. Chronoamperometry (−200 mV vs. SHE) of a carbon paper A. calcoaceticus biocathode (9 cm2 , pH 7.0, aerated) supplied with different concentrations of exogenous PQQ: the arrows indicate PQQ injection points (1.5 M per injection). The increase in catalytic current is roughly proportional to the concentration of added PQQ.
The development of catalytic current of oxygen reduction by S. putrefaciens can be followed in Fig. 2. The longer time required for the development of catalytic current compared to A. calcoaceticus is attributed to large amounts of carbon storage by S. putrefaciens (as revealed by 72 h long anodic current production by washed Shewanella cell in anodic experiments, data not shown). After 24 h of chronoamperometry at −200 mV SHE, cyclic voltammetry revealed a catalytic wave with onset at slightly less than +0.2 V SHE (Fig. 6). In the case of Shewanella, no peaks could be observed under anaerobic conditions, suggesting that possibly the last step of electron transfer is not carried out by a reversible redox compound. To determine the nature of the electron transfer, Shewanella cells were harvested from the bioelectrochemical reactor, washed twice with phosphate buffer saline, resuspended in phosphate buffer and immediately supplied to a pyrolytic graphite felt electrode with high surface area and activity for quick determination of the electrochemical response. In air-saturated conditions, a catalytic current with similar onset was observed (inset of Fig. 6), suggesting that redox compounds located on the external side of the outer membrane may be responsible for the last step of electron transfer in this case. The onset of the catalytic current is indeed close to the potential of some of the outer membrane cytochromes previously extracted from S. putrefaciens, including OmcA and
S. Freguia et al. / Electrochimica Acta 55 (2010) 813–818
Fig. 7. Close up of the onset of catalytic current as observed by cyclic voltammetry of carbon paper cathodes (9 cm2 ) at pH 7.0 and aerated conditions. Dotted line: abiotic control; solid line: S. putrefaciens biocathode; dashed line: abiotic cathode supplemented with 1 M hemin. Scan rate: 10 mV s−1 .
MtrC [30]. However, the filtered supernatant of the harvested cells exhibited exactly the same cyclic voltammetry shape and magnitude (see inset of Fig. 6), indicating that the compounds present on the surface of Shewanella’s outer membrane may also be released into the surrounding medium, where they may be able to carry out an abiotic catalytic effect similar to that of free hemin. A comparison of the onsets of the catalytic currents by Shewanella cells and hemin is shown in Fig. 7, revealing identical onsets for the two cases. The catalytic effect by the filtered supernatant may also be due to hemes released due to cell lysis. 4. Discussion Despite the possibility that oxygen reduction might occur based on non-biological pathways such as the ones described above (quinone autoxidation and direct electrochemistry of hemes), pathways that do require live bacteria are also present. The results obtained for A. calcoaceticus suggest the involvement of free PQQ in the catalysis. While PQQ autoxidation is not happening at significant rates, when the cofactor is supplied to the Acinetobacter biocathode, it is able to enhance the rate of cathodic oxygen reduction. Thus, there must be a pathway that involves the bacterium directly as a catalyst. Even though PQQ is normally bound to glucose dehydrogenase as a cofactor in Acinetobacter [19], pathways in which PQQ shuttles electrons between the soluble enzyme and an electrode have been previously proposed [31]. As the natural electron acceptor of PQQ is cytochrome c [19], it is likely that once PQQ is reduced by the cathode, electrons are transferred to cytochrome c. Finally cytochrome oxidases may complete the electron transfer to dioxygen, which is reduced to water. If we accept that PQQ is shuttling electrons between a cathode and the membranebound electron transfer chain, this proposed pathway becomes an easy possibility, which moreover would provide the bacteria with the important advantage that due to proton translocation during cytochrome c oxidation by complex IV, ATP may be produced through this reaction. In this case, bacteria would effectively use a cathode as insoluble electron donor for energy generation. While the results of this preliminary work suggest that this may be the case, proving that this indeed happens would require testing with cytochrome c and PQQ-dependent glucose dehydrogenase deletion mutants. However, in support of this theory, recent findings that bacteria can reduce nitrate (NO3 − ) to nitrite or N2 using a cathode as electron donor also point in this direction [32,33]. As nitrate reduction is not subject to enzyme-free reactions like O2 , it is most likely that in the case of nitrate reduction bacteria are exploiting
817
the electron transfer chain to deliver electrons from a cathode to the terminal electron acceptor. In the case of S. putrefaciens, membrane attached compounds appear to generate the catalytic current. As the bacterium is well known to localize c-type cytochromes on the external surface of the outer membrane, it is likely that cytochromes or cytochromecontaining nanowires are interacting directly with the cathode. However, in this case the electron transfer chain may be partially by-passed as oxygen may spontaneously react with hemes on the outer surface of the bacterium. The onset of oxygen reduction current with the bacteria tested in this study is at most +0.1 V vs. SHE. In two independent mixed culture studies [17,18], onsets as high as +0.45 V were obtained. More work needs to be done to isolate the bacteria responsible for such high onset of catalytic current, as well as to identify the bacterial redox compounds that enable such extracellular electron transfer. While enzymatic biocathodes can certainly achieve lower overpotentials with onsets of catalytic current as high as +0.75 V vs. SHE, the advantages of a self-renewable bacterial catalyst are numerous, justifying further research in this direction. 5. Conclusions The catalysis of oxygen reduction at microbial biocathodes may occur via several parallel pathways. Some of these pathways may be linked to the metabolic activity of the bacteria, while others are a side effect of bacterial excretion of redox active compounds which have the ability to chemically react with oxygen. Among redox active metabolites which are normally produced and excreted by bacteria, quinones in particular undergo the phenomenon of autoxidation, reducing O2 to O2 •− in solution and subsequently getting reduced at the cathode surface. Free hemes which may be excreted directly or exist in solution as lysis products have instead the ability to directly reduce O2 to H2 O2 at the cathode surface. The extracellular electron pathways which require the metabolic activity of the bacteria directly are likely to involve the electrical interaction between the cathode and the bacteria’s electron transfer chain. This interaction may be carried out via shuttling of excreted quinones, or via direct contact by redox active compounds (such as c-type cytochromes) located on the outer membrane of the bacteria. Acknowledgement The first author was supported by the Japan Society for the Promotion of Science (PE 08026). References [1] C.W. Lee, H.B. Gray, F.C. Anson, B.G. Malmstrom, J. Electroanal. Chem. (1984) 289. [2] M.R. Tarasevich, A.I. Yaropolov, V.A. Bogdanovskaya, S.D. Varfolomeev, Bioelectrochem. Bioenerg. 6 (1979) 393. [3] G.T.R. Kim, H.H. Palmore, J. Electroanal. Chem. 464 (1999) 110. [4] S. Tsujimura, Y. Miura, K. Kano, Electrochim. Acta 53 (2008) 5716. [5] S. Tsujimura, B. Tatsumi, J. Ogawa, S. Shimizu, K. Kano, T. Ikeda, J. Electroanal. Chem. 496 (2001) 69. [6] S. Tsujimura, Y. Kamitaka, K. Kano, Fuel Cells 7 (2007) 463. [7] C. Vaz-Dominguez, S. Campuzano, O. Rudiger, M. Pita, M. Gorbacheva, S. Shleev, V.M. Fernandez, A.L. De Lacey, Biosens. Bioelectron. 24 (2008) 531. [8] K. Rabaey, N. Boon, S.D. Siciliano, M. Verhaege, W. Verstraete, Appl. Environ. Microbiol. 70 (2004) 5373. [9] D.E. Holmes, S.K. Chaudhuri, K.P. Nevin, T. Mehta, B.A. Methe, A. Liu, J.E. Ward, T.L. Woodard, J. Webster, D.R. Lovley, Environ. Microbiol. 8 (2006) 1805. [10] B.H. Kim, T. Ikeda, H.S. Park, H.J. Kim, M.S. Hyun, K. Kano, K. Takagi, H. Tatsumi, Biotechnol. Tech. 13 (1999) 475. [11] H. Richter, K.P. Nevin, H.F. Jia, D.A. Lowy, D.R. Lovley, L.M. Tender, Energy Environ. Sci. 2 (2009) 506.
818
S. Freguia et al. / Electrochimica Acta 55 (2010) 813–818
[12] G. Reguera, K.D. McCarthy, T. Mehta, J.S. Nicoll, M.T. Tuominen, D.R. Lovley, Nature 435 (2005) 1098. [13] A. Bergel, D. Feron, A. Mollica, Electrochem. Commun. 7 (2005) 900. [14] P. Clauwaert, D. Van der Ha, N. Boon, K. Verbeken, M. Verhaege, K. Rabaey, W. Verstraete, Environ. Sci. Technol. 41 (2007) 7564. [15] S. Freguia, K. Rabaey, Z. Yuan, J. Keller, Water Res. 42 (2008) 1387. [16] V. Scotto, R. Dicintio, G. Marcenaro, Corros. Sci. 25 (1985) 185. [17] K. Rabaey, S.T. Read, P. Clauwaert, S. Freguia, P.L. Bond, L.L. Blackall, J. Keller, ISME J. 2 (2008) 519. [18] A. Aldrovandi, E. Marsili, L. Stante, P. Paganin, S. Tabacchioni, A. Giordano, Bioresour. Technol. 100 (2009) 3252. [19] K. Matsushita, H. Toyama, M. Yamada, O. Adachi, Appl. Microbiol. Biotechnol. 58 (2002) 13. [20] C.R. Myers, J.M. Myers, J. Bacteriol. 174 (1992) 3429. [21] E. Marsili, D.B. Baron, I.D. Shikhare, D. Coursolle, J.A. Gralnick, D.R. Bond, Proc. Natl. Acad. Sci. U.S.A. 105 (2008) 3968. [22] H.J. Kim, H.S. Park, M.S. Hyun, I.S. Chang, M. Kim, B.H. Kim, Enzyme Microb. Technol. 30 (2002) 145.
[23] Y.A. Gorby, S. Yanina, J.S. McLean, K.M. Rosso, D. Moyles, A. Dohnalkova, T.J. Beveridge, I.S. Chang, B.H. Kim, K.S. Kim, D.E. Culley, S.B. Reed, M.F. Romine, D.A. Saffarini, E.A. Hill, L. Shi, D.A. Elias, D.W. Kennedy, G. Pinchuk, K. Watanabe, S. Ishii, B. Logan, K.H. Nealson, J.K. Fredrickson, Proc. Natl. Acad. Sci. U.S.A. 103 (2006) 11358. [24] H. Tatsumi, H. Nakase, K. Kano, T. Ikeda, J. Electroanal. Chem. 443 (1998) 236. [25] H. Iken, L. Etcheverry, A. Bergel, R. Basseguy, Electrochim. Acta 54 (2008) 60. [26] Bergel v, A. Lai, J. Electroanal. Chem. 494 (2000) 30. [27] S. Freguia, M. Masuda, S. Tsujimura, K. Kano, Bioelectrochemistry 76 (2009) 14. [28] D. Compton, R.G. Menshykau, Electroanalysis 20 (2008) 2387. [29] S. Itoh, H. Kawakami, S. Fukuzumi, Chem. Commun. (1997) 29. [30] M. Firer-Sherwood, G.S. Pulcu, S.J. Elliott, J. Biol. Inorg. Chem. 13 (2008) 849. [31] V. Laurinavicius, J. Razumiene, A. Ramanavicius, A.D. Ryabov, Biosens. Bioelectron. 20 (2004) 1217. [32] K.B. Gregory, D.R. Bond, D.R. Lovley, Environ. Microbiol. 6 (2004) 596. [33] B. Virdis, K. Rabaey, Z. Yuan, J. Keller, Water Res. 42 (2008) 3013.