Catalytic response of microbial biofilms grown under fixed anode

Chemical Engineering Journal 230 (2013) 532–536

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Catalytic response of microbial biofilms grown under fixed anode potentials depends on electrochemical cell configuration Amit Kumar a,b,c,⇑, Alma Siggins b,d, Krishna Katuri b, Therese Mahony b,d, Vincent O’Flaherty b,d, Piet Lens c, Dónal Leech a,b,⇑ a

Biomolecular Electronics Research Laboratory, School of Chemistry, National University of Ireland Galway, Ireland Ryan Institute, National University of Ireland Galway, Ireland Department of Environmental Engineering and Water Technology, UNESCO-IHE, Delft, The Netherlands d School of Natural Sciences, National University of Ireland Galway, Ireland b c

h i g h l i g h t s

g r a p h i c a l a b s t r a c t

Microbial biofilms show different

catalytic response at different anode potentials. Membrane-less bioelectrochemical cell poised at +0.2 V vs. Ag/AgCl showed maximum current density of >5000 mA/m2. Membrane separated bioelectrochemical cell poised at 0.3 V vs. Ag/AgCl showed current density of 4000 mA/m2. Preliminary microbial analysis shows differences in both configurations.

a r t i c l e

i n f o

Article history: Received 29 December 2012 Received in revised form 22 May 2013 Accepted 16 June 2013 Available online 27 June 2013 Keywords: Bioelectrochemistry Microbial biofilm Fuel cell Applied anode potential Bioenergy Bioelectrochemical reactor configuration

a b s t r a c t In microbial electrochemical cells the anode potential can vary over a wide range, which alters the thermodynamic energy available for bacterial-electrode electron exchange (termed electroactive bacteria). We investigated how anode potential affected the microbial catalytic response of the electroactive biofilm. Microbial biofilms induced to grow on graphite electrodes by application of a fixed applied anode potential in membrane-separated and membrane-less electrochemical cells show differences in electrocatalytic response. Maximum current density is obtained using +0.2 V vs. Ag/AgCl to induce biofilm growth in membrane-less cells, in contrast to a maximum achieved at lower applied potentials in a membrane-separated electrochemical cell configuration. This insight into differences in optimal applied potentials based on cell configuration can play an important role in selection of parameters required for microbial fuel cells and bio-electrochemical systems. Ó 2013 Elsevier B.V. All rights reserved.

1. Introduction In microbial electrogenesis bacteria oxidize substrates to generate electricity by transferring electrons to a solid electrode, leading ⇑ Corresponding authors. Address: Biomolecular Electronics Research Laboratory, School of Chemistry, National University of Ireland Galway, Ireland. E-mail addresses: [email protected] (A. Kumar), donal.leech@ nuigalway.ie (D. Leech). 1385-8947/$ - see front matter Ó 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.cej.2013.06.044

to potential for electricity generation in microbial fuel cells (MFC) or to offset the potential required for hydrogen, and other valueadded chemical, production at cathodes in bio-electrochemical systems (BES) [1,2]. BES operating conditions affect growth kinetics and metabolism of the microbial community (Fig. 1). In microbial fuel cells, with current flow between an anode and a cathode across a fixed resistance load, the potential imposed on the anode is difficult to control, because of variations in anode potential as a function of mass transport to, catalytic activity at, and current flow

A. Kumar et al. / Chemical Engineering Journal 230 (2013) 532–536

between, anode and cathode. Growth of microbial biofilms can however be induced by precise control of anode potential vs. a reference electrode using a potentiostat. The electron transfer capabilities of microbial films induced to grow by imposition of specific anode potentials can be thus explored in fundamental studies of electroactive biofilms on electrodes. However, results comparing applied anode potentials show a range of response in terms of effect of potential on fuel cell power and/or current density [3–8], as reviewed recently by Kumar et al. [4]. There are few studies comparing current densities achieved using a fixed resistance load MFC to induce microbial biofilm growth on anodes to those using fixed anode potentials. Wang et al. [9] compare biofilm growth under an applied anode potential of +0.4 V vs. Ag/AgCl to that across a fixed 1000 X resistance load between anode and a ferricyanide-reducing cathode, separated by an Ultrex cation exchange membrane. The application of a fixed anode potential produced a reproducible current in fewer cycles, and shorter time, compared to that using the fixed resistance load cell. Other studies on BESs using either individual applied anode potentials (different values), or a single applied potential, show variable results in terms of current or power generation, as reviewed recently [4]. There are however no reports to our knowledge comparing electrocatalytic properties of anode potential-induced biofilms as a function of the electrochemical cell configuration (i.e., in membrane-separated vs. membrane-less electrochemical cells). When MFCs and BESs are operated, anode potentials measured during peak power generation show typical values around 0.40 to 0.48 V vs. Ag/AgCl for mixed cultures oxidizing acetate [4]. Applied anode potentials more positive than this therefore provide a driving force for microbially catalyzed oxidation of acetate in biofilms on anodes. We probe the effect of 0.3 V, 0.2 V and +0.2 V vs. Ag/AgCl applied anode potentials on biofilm performance, as a function of electrochemical cell

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configuration, using anaerobic sludge as a mixed culture inoculum and acetate as feed. 2. Experimental section 2.1. Reactor configuration and operation Experiments were conducted in either membrane-less or membrane-separated electrochemical cell configuration. The membrane-less configuration was a single borosilicate glass cell containing a Ag/AgCl (3 M NaCl) reference electrode, and both anode and cathode electrodes. For the membrane-separated configuration, the anodic and cathodic borosilicate glass half-cells were Ò separated by a 12 cm2 Nafion 117 proton exchange membrane (Sigma, Ireland), with the reference electrode in the anode halfcell. In both configurations the anode was a 3.9 mm diameter graphite rod electrode (5 cm2 geometric exposed area) and the cathode was a carbon cloth (24 cm2, E-TEK, nonwet-proof). The membrane-less cell experiments used 150 mL of feed solution, purged with oxygen free nitrogen gas to maintain an anaerobic environment, whilst for membrane-separated cell experiments both half-cell compartments were fed with anaerobic cell culture medium solution (each half-cell 150 mL), to permit comparative studies where the only variable is the cell configuration. Growth medium containing 20 mM acetate was prepared according to culture centre protocol (http://www.dsmz.de, medium No. 826). All bioelectrochemical cells were operated in batch mode initially (0–144 h) and then switched to continuous feed (1 day hydraulic retention time, 20 mM acetate medium, up to 660 h) mode, at 30 °C. In all bioelectrochemical cells, anodes were polarized independently (a Ag/AgCl reference electrode for each channel) by a multi-channel Uniscan PG580RM potentiostat. Electrochemical cells were subjected to real-time electrochemical measurements

Fig. 1. Impact of various factors on performance of bioelectrochemical processes.

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as described by Katuri et al. [7]. All potentials are measured and quoted vs. a Ag/AgCl (3 M NaCl) reference electrode unless otherwise stated. 2.2. Preparation of graphite rod electrodes Custom built electrodes were made by shrouding graphite rods (0.39 cm £, Graphite store NC001300 product) in heat-shrink tubing (Alphawire, UK) and establishing an electrical connection at the rear with a copper rod (0.3 cm £; Farnell electronics, Ireland) and silver epoxy adhesive (Radionics, Ireland). Electrodes were sterilized by boiling in 0.1 M H2SO4 for 15 min, followed by storage in absolute ethanol. 2.3. Inoculum Granular anaerobic sludge from a wastewater bioreactor (Carberry Milk Products, Ireland) was crushed and concentrated (5000 g, 10 min at 20 °C), washed and re-suspended in a ratio of 1:20 v/v in sterile degassed phosphate buffer (0.1 M, pH 7.0). 2.4. Microbial diversity and similarity analysis Following cessation of operation of electrochemical experiments (660 h) the electrodes were removed from the cells, sonicated for 10 s (MSE Soniprep 150) at an amplitude of 22 lm and DNA extracted from 100 lL of the cell suspension using an automated nucleic acid extractor (Magtration 12GC, PSS Co., Chiba Japan) with extracted DNA eluted in 100 lL Tris–HCl buffer (pH 8.0) and stored at 20 °C. Denaturing gradient gel electrophoresis (DGGE) was performed as described in Siggins et al. [10]. Briefly, initial PCR amplification used the primer set 341F (50 -CCT ACG GGA GGC AGC AG-30 ) and 517R (50 -ATT ACC GCG GCT GCT GG-30 ) [11] with 40-base pair GC clamp on the 50 terminus of the forward primer. Touchdown PCR was of initial denaturation at 94 °C for 2 min, followed by denaturation at 94 °C for 30 s, annealing of primers (65–55 °C; 1 cycle at 1 °C increments; 20 cycles at 55 °C) for 30 s and extension at 72 °C for 30 s, followed by a final extension at 72 °C for 10 min with PCR products electrophoresed on 2% agarose gels. Statistical analysis of DGGE gels using TotalLab software (Phoretix, Newcastle, United Kingdom) used scoring of presence or absence of bands with ‘‘1’’ or ‘‘0’’, respectively, with binary matrices used to calculate unweighted pair-group method using arithmetic averages (UPGMA) dendrograms, based on Bray–Curtis similarity co-efficients (MVSP Version 3; Kovach Computing Services). Bands (23 in total) were excised and suspended in 200 lL of sterile water to elute DNA which was then PCR amplified and cloned using TOPOÒ TA (Invitrogen) and selected DNA fragments were sequenced (MWG). The Mega5 software was used to align sequences with 16S rRNA gene sequences retrieved from the BLASTn database, and to carry out phylogenetic analysis. As the resulting partial 16S rDNA gene sequences were 5000 mA m2) under steady-state continuous feed conditions, compared to that observed for biofilms induced to grow under more negative applied anode potentials (0.3 V vs. Ag/AgCl produced 65% similarity for samples from anodes in membrane-separated configuration, with that from the anode at +0.2 V vs. Ag/AgCl showing greatest divergence from others for both configurations (Fig. S1, ESI). Biofilm samples from anodes in membrane-separated (dual chamber) configuration also showed a different community composition to that of biofilm samples from anodes and cathodes in membrane-less (single chamber) configuration (Fig. S2, ESI).

4. Conclusions, outlook and implications Linking anode potential to evolution of microbial biofilm current and/or power generation and microbial diversity information has been an issue of debate over the past decade. Our preliminary results indicate that consideration of the electrochemical

cell configuration, with or without separating membrane, is required in any attempt to select an applied anode potential or to alter microbial activity in such systems. Among membrane-less bioelectrochemical cells, the electrochemical cell poised at the highest potentials produced the highest steady-state electrocatalytic current densities for acetate oxidation in a cyclic voltammogram, whilst the electrode at the lowest potential produced a maximum of 1500 mA/m2. However, among the membrane separated bioelectrochemical cells (H-cell), the electrochemical cells poised at 0.3 and 0.2 V vs. Ag/AgCl potentials developed higher steady-state current densities (4000 and 6000 mA/m2, respectively) than anodes poised at higher potentials (2000 mA/m2 for +0.2 V vs. Ag/AgCl) in their cyclic voltammetric curves. The anode community structures of the three single chambered bioelectrochemical cells were P76% similar, with the anode from the bioelectrochemical cell poised at +0.2 V vs. Ag/AgCl showing the greatest divergence. The anode biofilm samples of the three membrane-separated bioelectrochemical cells demonstrated >65% similarity to each other. Phylogenetic analysis of the sequences obtained from the membrane-separated bioelectrochemical cell biomass samples showed a different community composition than that of the membrane-less bioelectrochemical cell biomass. Clearly further investigations are needed to better understand this link. These should explore the role of reactions occurring as a result of

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electrochemical (and bioelectrochemical) reductions at cathodes in membrane-less and membrane-separated configurations. Furthermore, the effect due to the introduction of oxidants (e.g., oxygen), as a recent study on MFCs reported differences in microbial diversity due to oxygen exposure [26], or production of biogas (e.g., methane or hydrogen) on biofilm production and activity in these systems warrants investigation. Such an approach may improve understanding of mechanism of reactions involved. In addition, insight provided by this study could be useful for selecting appropriate microorganism(s) and elucidating the role of different electroactive processes and electron transfer mechanisms in BES. Acknowledgments Authors acknowledge funding from FP 7 People Programme of European Commission, Marie Curie Intra European Fellowship for Career Development (Grant A/6342 – PIEF-GA-2009-237181) and Science Foundation Ireland (Charles Parsons Energy Research Award – 06/CP/E006). Appendix A. Supplementary material Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.cej.2013.06.044. References [1] D.R. Lovley, Powering microbes with electricity: direct electron transfer from electrodes to microbes, Environmental Microbiology Reports 3 (2011) 27–35. [2] A.E. Franks, K.P. Nevin, R.H. Glaven, D.R. Lovley, A novel approach for spatial analysis of global gene expression within a Geobacter sulfurreducens currentproducing biofilm, ISME Journal 4 (2010) 509–519. [3] P. Alterman, S. Freguia, J. Keller, W. Verstraete, K. Rabaey, The anode potential regulates bacterial activity in microbial fuel cells, Applied Microbial Biotechnology 78 (2008) 409–418. [4] A. Kumar, K. Katuri, P. Lens, D. Leech, Does bioelectrochemical cell configuration and anode potential affect biofilm response?, Biochemical Society Transactions 40 (2012) 1308–1314 [5] R.C. Wagner, D.F. Call, B.E. Logan, Optimal set anode potentials vary in bioelectrochemical systems, Environmental Science Technology 44 (2010) 6036–6041. [6] C.I. Torres, B.R. Krajmalnik, P. Parameswaran, A.K. Marcus, G. Wanger, Y.A. Gorby, B.E. Rittmann, Selecting anode-respiring bacteria based on anode potential: phylogenetic, electrochemical, and microscopic characterization, Environmental Science Technology 43 (2009) 9519–9524. [7] K.P. Katuri, P. Kavanagh, S. Rengaraj, D. Leech, Geobacter sulfurreducens biofilms developed under different growth conditions on glassy carbon electrodes: insights using cyclic voltammetry, Chemical Communications 46 (2010) 475847–475860. [8] C.I. Torres, A.K. Marcus, P. Parameswaran, B.E. Rittmann, Kinetic experiments for evaluating the Nernst–Monod model for anode-respiring bacteria (ARB) in a biofilm anode, Environmental Science Technology 42 (2008) 6593–6597.

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