APPLIED AND ENVIRONMENTAL MICROBIOLOGY, Dec. 2008, p. 7329–7337 Vol. 74, No. 230099-2240/08/$08.00�0 doi:10.1128/AEM.00177-08Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Microbial Biofilm Voltammetry: Direct Electrochemical Characterizationof Catalytic Electrode-Attached Biofilms�†

Enrico Marsili,1 Janet B. Rollefson,2 Daniel B. Baron,1Raymond M. Hozalski,3 and Daniel R. Bond1,4*

BioTechnology Institute, University of Minnesota, St. Paul, Minnesota 551081; Department of Biochemistry, Molecular Biology andBiophysics, University of Minnesota, Minneapolis, Minnesota 554552; Department of Civil Engineering, University of Minnesota,

Minneapolis, Minnesota 554553; and Department of Microbiology, University of Minnesota, Minneapolis, Minnesota 554554

Received 18 January 2008/Accepted 21 September 2008

While electrochemical characterization of enzymes immobilized on electrodes has become common, there isstill a need for reliable quantitative methods for study of electron transfer between living cells and conductivesurfaces. This work describes growth of thin (<20 �m) Geobacter sulfurreducens biofilms on polished glassycarbon electrodes, using stirred three-electrode anaerobic bioreactors controlled by potentiostats and nonde-structive voltammetry techniques for characterization of viable biofilms. Routine in vivo analysis of electrontransfer between bacterial cells and electrodes was performed, providing insight into the main redox-activespecies participating in electron transfer to electrodes. At low scan rates, cyclic voltammetry revealed catalyticelectron transfer between cells and the electrode, similar to what has been observed for pure enzymes attachedto electrodes under continuous turnover conditions. Differential pulse voltammetry and electrochemical im-pedance spectroscopy also revealed features that were consistent with electron transfer being mediated by anadsorbed catalyst. Multiple redox-active species were detected, revealing complexity at the outer surfaces ofthis bacterium. These techniques provide the basis for cataloging quantifiable, defined electron transferphenotypes as a function of potential, electrode material, growth phase, and culture conditions and provide aframework for comparisons with other species or communities.

Anaerobic metal-reducing Geobacteraceae catalyze thetransfer of electrons from reduced electron donors, such asacetate, to oxidized electron acceptors such as Fe(III). Whileelectron transport is confined to the inner membrane of mostrespiratory bacteria (66), Fe(III) and Mn(IV) exist primarily inthe environment as insoluble precipitates at circumneutral pH.To reduce minerals of various crystallinity, redox potential,and surface charge, metal-reducing bacteria must possess acomplete mechanism for transporting electrons across the in-ner membrane, periplasm, outer membrane, and outer sur-faces (23, 38, 48, 54, 57, 58). Soluble redox-active shuttles orchelators can facilitate respiration beyond the outer membranein some organisms (12, 34, 55, 56, 63), but no evidence for thesecretion of such a compound has been observed in Geobacter-aceae.

This electron transfer capability has recently been utilized indevices known as microbial fuel cells (MFCs), where elec-trodes serve as electron acceptors to support growth (11, 18,35, 36, 62). However, variations in electrode surfaces, operat-ing temperatures, electron donor concentrations, and cultureconditions limit comparison between electrode-reducing bac-teria. In addition, MFCs aim to maximize electrical powerproduction per unit volume, using large anodes consisting ofcarbon felt (19, 39, 50) or carbon paper (43) in small reactors

(with surface/volume ratios between 25 and 640) (19, 25, 29,43). In such systems, the actual surfaces accessible to bacteriaare not known, multiple “dead zones” or diffusion-limited re-gions exist (19), high-surface-area electrodes increase capaci-tive current and reduce the sensitivity of analyses (53, 68), andirregular surfaces prevent mathematical modeling or imagingof attached biomass (22).

Electrochemical techniques typically used to study purifiedredox proteins (3, 4, 7) and cell extracts (26, 45) could likely beadapted for the study of metal-reducing bacteria. While pro-tein voltammetry is limited by the success of purification andimmobilization of enzymes (17, 49), metal-reducing bacteriarepresent a self-assembling multienzyme system that naturallyinterfaces with electrodes. In this report, we describe proce-dures for analysis of G. sulfurreducens biofilms on small carbonelectrodes of defined roughness in potentiostat-controlledthree-electrode cells. Several direct current and alternatingcurrent techniques could be applied to the characterization ofthe biofilm-electrode interface and have produced insights intoelectron transfer mechanisms between living cells and elec-trodes. Application of these techniques in routine character-ization of other living bacteria and biofilm communities willallow quantitative comparisons between electron transfer ratesand mechanisms used by these organisms.

MATERIALS AND METHODS

Bacterial strain and culture media. G. sulfurreducens strain PCA (ATCC51573) was subcultured in our laboratory at 30°C using a vitamin-free anaerobicmedium containing the following per liter: 0.38 g KCl, 0.2 g NH4Cl, 0.069 gNaH2PO4 � H2O, 0.04 g CaCl2 � 2H2O, 0.2 g MgSO4 � 7H2O, and 10 ml of amineral mix [containing 0.1 g MnCl2 � 4H2O, 0.3 g FeSO4 � 7H2O, 0.17 gCoCl2 � 6H2O, 0.1 g ZnCl2, 0.04 g CuSO4 � 5H2O, 0.005 g AlK(SO4)2 � 12H2O,

* Corresponding author. Mailing address: BioTechnology Institute,University of Minnesota, 140 Gortner Laboratory, 1479 Gortner Ave.,St. Paul, MN 55108. Phone: (612) 624-8619. Fax: (612) 625-1700.E-mail: dbond@umn.edu.

† Supplemental material for this article may be found at http://aem.asm.org/.

� Published ahead of print on 10 October 2008.

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0.005 g H3BO3, 0.09 g Na2MoO4, 0.12 g NiCl2, 0.02 g NaWO4 � 2H2O, and 0.10 gNa2SeO4 per 1 liter]. Acetate was provided as an electron donor at 20 mM. Allmedia were adjusted to pH 6.8 prior to the addition of 2 g/liter NaHCO3 andwere flushed with oxygen-free N2-CO2 (80:20 [vol/vol]) prior to sealing with butylrubber stoppers and autoclaving. All experiments were initiated by inoculatingwith 40% (vol/vol) of cells within 3 h of reaching maximum optical density (�0.6)in the medium described.

Electrode preparation. Glassy carbon (Toko America, New York, NY) wasmachine cut into 2- by 0.5- by 0.1-cm electrodes. Freshly cut glassy carbonelectrodes were polished using aluminum oxide-silicon carbide sandpaper with agrit designation of between P400 and P4000 (3M, Minneapolis, MN), as indi-cated in the ISO/FEPA standard (http://www.fepa-abrasives.com/). Mirror-pol-ished electrode surfaces were obtained with 0.05-�m alumina powder (CH In-struments, Austin, TX). Polished electrodes were sonicated to remove debris,soaked overnight in 1 N HCl to remove metals and other contaminants, washedtwice with acetone and deionized water to remove organic substances, and storedin deionized water. After each experiment, electrode surfaces were cleaned withan additional 1 N NaOH treatment (to remove biomass), and the entire surfacewas refreshed through sandpaper polishing and cleaning as described above toremove immobilized electron transfer agents. These working electrodes wereattached to 0.1-mm Pt wires via miniature nylon screws that ensured electricalcontact throughout the experiment.

Electrode cell assembly. Platinum wires from the working electrode wereinserted into heat-pulled 3-mm glass capillary tubes (Kimble, Vineland, NJ) andsoldered inside the capillary to copper wires. Counter electrodes consisted of a0.1-mm-diameter Pt wire (Sigma-Aldrich, St. Louis, MO) that was also insertedinto a 3-mm glass capillary and soldered to a copper wire. The resistance of eachelectrode assembly was measured, and electrodes with a total resistance of higherthan 0.5 � were discarded. Reference electrodes were connected to bioreactorsvia a salt bridge assembled from a 3-mm glass capillary and a 3-mm Vycor frit(BioAnalytical Systems, West Lafayette, IN). Electrode capillaries were inserted

through ports in a custom-made Teflon lid that was sealed with an O ring gasket.This lid fit onto a 20-ml conical electrochemical cell (BioAnalytical Systems,West Lafayette, IN), which had been previously washed in 3 N HNO3 (Fig. 1A).

After the addition of a small magnetic stir bar, the cell was autoclaved for 15min. Following autoclaving, the salt bridge was filled with 0.1 M Na2SO4 in 1%agar. A saturated calomel reference electrode (Fisher Scientific, Pittsburg, PA)was placed at the top of this agar layer and covered in additional Na2SO4 toensure electrical contact (60). The reactors were placed in an in-house-builtwater bath to maintain cells at 30°C (which introduced less electrical noise thanlarge circulators, incubators, or temperature-controlled rooms) (Fig. 1B). Tomaintain the strict anaerobic conditions required by bacteria, all reactors wereoperated under a constant flow of sterile humidified N2-CO2 (80:20 [vol/vol]),which had been passed over a heated copper column to remove trace oxygen.Each reactor was located above an independent magnetic stirring unit.

Autoclaved bioreactors flushed free of oxygen, filled with sterile growth me-dium, and incubated at 30°C were analyzed before each experiment to verifyanaerobicity and the absence of redox-active species. Electrochemical cells show-ing residual peaks in differential pulse voltammetry (DPV), anodic current incyclic voltammetry (CV), or baseline noise were discarded as having possibleelectrode cleanliness or connection noise issues. These autoclaved, verified bio-reactors were then used for growth of G. sulfurreducens cultures.

A typical bioreactor was inoculated with 40% (vol/vol) of stationary phase G.sulfurreducens cells. After inoculation, a potential step of 0.24 V versus thestandard hydrogen electrode (SHE) was applied and the reactors were incubateduntil the current stabilized (�72 h). The potential step was �200 mV higher thanthe midpoint potential, sustaining a half-maximal anodic current (as determinedby preliminary CV; see below) (9).

Electrochemical instrumentation. A 16-channel potentiostat (VMP, Bio-Logic, Knoxville, TN) was connected to the three-electrode cells described above(Fig. 1B). Software from the same producer (EC-Lab v9.41) was used to runsimultaneous multitechnique electrochemistry routines, which include CV, DPV,

FIG. 1. Cartoon of three-electrode bioelectroreactor (A) and connection scheme to multichannel potentiostat (B).

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and chronoamperometry (CA). Scan rates higher than 100 mV/s and all electro-chemical impedance spectroscopy (EIS) experiments were performed using aGamry PCI4 Femtostat (Gamry Instruments, Warminster, PA), and EIS datafitting was performed with E-Chem Analyst v5.1. Postacquisition analysis of CVdata was performed with the software Utilities for Data Analysis (UTILS), kindlyprovided by D. Heering (v1.0; University of Delft, The Netherlands [http://www.tudelft.nl/]). All measurements, with the exception of CA, were performed insuccession without stirring enabled. The parameters for DPV were as follows: Einitial

(Ei) � �0.558 V versus SHE and Efinal (Ef) � 0.242 V versus SHE; pulse height,50 mV; pulse width, 300 ms; step height, 2 mV; step time, 500 ms; scan rate, 4mV/s; current averaged over the last 80% of the step (1 s, 12 points); accumu-lation time, 5 s. The parameters for CV were as follows: equilibrium time, 5 s;scan rate, 1 mV/s; Ei � �0.558 V versus SHE; Ef � 0.242 V versus SHE;current averaged over the whole step (1 s, 10 points). For CA, the parameterswere Eapplied (E) � 0.242 V versus SHE.

EIS was conducted while maintaining a direct current voltage between theworking electrode and the reference electrode (potentiostatic EIS) as electrontransfer processes by Geobacter isolates were observed to be voltage dependent(62). EIS was performed at four potentials, 0.04, �0.06, �0.16, and �0.26 Vversus SHE, with a perturbation amplitude of 10 mV. The frequency was variedfrom 105 to 0.01 Hz, as most biological charge transfer phenomena are observedin this range (10). Data were then fitted with a simple empirical model (62), usinga constant phase element to account for irregularities in the morphology andchemistry of the glassy carbon surface. EIS was typically performed only at theend of incubations as extensive characterization required electrodes to be held atmultiple potentials for long periods of time.

Confocal and SEM analyses. A S3500N scanning electron microscope (SEM)(Hitachi, Japan) was used to image freshly polished (SEM) electrodes. Afterbeing thoroughly cleaned with deionized water, 2- by 0.5- by 0.1-cm graphiteelectrodes freshly polished with P400 and P4000 sandpapers were air dried andfixed through graphite tape to a sample stub. The sample stubs were placeddirectly into the SEM vacuum compartment and examined at an acceleratingvoltage of 5 kV.

A Nikon C1 spectral imaging confocal microscope (Nikon, Japan) with a �60lens was used to image biofilm-covered electrodes. Immediately following har-vesting, the biofilm-covered electrodes were gently washed in growth medium toremove planktonic cells and then stained with the LIVE/DEAD Baclight bacte-rial viability kit (Invitrogen Corp., Carlsbad, CA), incubated for 15 min in thedark, rinsed with growth medium, and placed on a microscope slide. The cov-erslip was supported by glass spacers thicker than the electrode to preventdamage to biofilm structure. Two laser wavelengths, 488 and 561 nm, were usedto excite the two stains.

RESULTS AND DISCUSSION

Colonization of electrodes. In MFCs, the half-cell potentialat the anode varies with the activity of the bacterial biofilm andthe chosen external resistance (37, 43). Potentiostat-controlledelectrodes at a sufficiently positive potential are equivalent toanodes in which the electron acceptor is nonlimiting, reducingthe technical complexity and simplifying the conceptual modelof electron transfer. However, at high oxidative potentials,conformational change, unfolding, and irreversible processcould alter the catalytic abilities of enzymes, decreasing theanodic (oxidation) current (27, 52). In all experiments de-scribed, the potential step used was 0.24 V versus SHE. Thispotential was chosen based on preliminary CV experiments,which showed that the oxidation current from cells wasindependent of applied potential in the range of 0.1 to 0.3 Vversus SHE.

After inoculation, the potential step was applied, and a rap-idly increasing anodic (oxidation) current was detected (Fig.2). As this rate of increase was many times higher than whatcould be explained by known growth rates of G. sulfurreducens,it was likely due to cell attachment to the electrode. Theattachment phase was most rapid when fumarate-limited cells(compared to mid-log-phase cells) were used as the inoculum,

consistent with reports linking acceptor limitation to the ex-pression of enzymes important to metal reduction (21, 23).This was followed by a growth phase, characterized by anexponential increase in current, which doubled at a rate typi-cally observed for G. sulfurreducens reducing Fe(III)-citrate orfumarate (doubling time, �7 h).

Within 72 h, electrodes reached a saturation phase charac-terized by a stable maximum electron transfer rate with astandard deviation between biological replicates of less than3% (n � 6). Confocal imaging of electrodes always revealeduniform thin films of �15 �m with few structural features suchas pillars (see Fig. S1 in the supplemental material). During theattachment and growth phases, electrodes were tested for theeffect of voltammetry analyses on the rate of growth or currentplateau. Repeated CV and DPV (see Materials and Methods)did not alter key midpoint potentials, colonization rates, orfinal current densities (Fig. 2).

In order to minimize contribution of other electron transferagents, we used a medium lacking redox-active species, such asvitamins (i.e., riboflavin), redox indicators (resazurin), andchemical reductants (cysteine and dithiothreitol) (47, 64). Inour recent study (47), we showed that flavin concentrations inthe nM range can be detected under the experimental condi-tions applied in the present work. When the medium in reac-tors was replaced, removing planktonic cells and any potentialmicrobially produced soluble electron transfer agent, the elec-tron transfer rate to electrodes remained unchanged. In addi-tion, removal of nitrilotriacetic acid-chelated minerals fromthe medium (once electrode colonization had occurred) didnot affect current production. These findings created stringentconditions for collection of baseline data in the absence ofmost common soluble redox-active compounds. A systematicexamination of the effects of commonly present redox species

FIG. 2. Characteristic growth curve for G. sulfurreducens inocu-lated into a bioreactor containing a 2.5-cm2 glassy carbon electrode(P400 grit polishing treatment). “Analysis” breaks indicate when CVand DPV were performed. “Medium change” breaks indicate whereplanktonic cells and culture medium were removed and replaced withfresh medium. Raw data are shown with no smoothing or noise-reduc-tion transformations.

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on electron transfer by Geobacter spp. will be the focus of afuture report.

The effect of microscale surface roughness on bacterial at-tachment and direct electron transfer has not been reported.Figure 3 shows the surface features of carbon electrodes pol-ished with different sandpapers. When G. sulfurreducens bio-films were grown on electrodes polished with increasing gritsize, the maximum current density increased over 100% for thesame geometric area (Fig. 3). Polishing with particles smallerthan �5 �m did not significantly reduce the maximum current.Therefore, even micrometer scale features have to be con-trolled to allow reproducibility and comparisons between cul-tures and electrodes in this and future studies.

CV. CV is the principal diagnostic method in protein filmvoltammetry, which studies immobilized enzymes on rotatingand stationary carbon electrodes. When in the presence of asuitable substrate and slowly cycled through a range of appliedpotentials at low scan rates, enzymes have time to undergomultiple turnovers at each sampled potential (5–7, 33). As scanrates increase, slow reactions may not have time to occurbefore the potential changes to the next step, and concentra-tion gradients representative of steady-state conditions maynot have time to establish. Thus, as the scan rate increases, thekinetics of interfacial electron transfer between redox pro-tein(s) and electrodes affects the voltammetric response more

strongly and can hide the kinetics of continuous enzymaticturnover. As the purpose of this study was to establish methodsfor measuring catalytic (turnover) kinetics in viable bacterialbiofilms, in the absence of prior knowledge regarding interfa-cial electron transfer rates, low scan rates represented the mostconservative experimental conditions. Parameters for the studyof nonturnover voltammetry at high scan rates will be dis-cussed in subsequent reports.

Application of CV to living bacteria required limiting thepotential range to prevent harmful oxidizing or reducing con-ditions, as well as selection of informative scan rates. While itis known that application of potentials can disrupt biofilmseither by producing hydrogen (28) or by inducing unfolding/oxidation of adsorbed proteins at oxidizing potentials (52), G.sulfurreducens biofilms were not harmed by exposure to poten-tials between �0.55 and 0.24 V versus SHE. For example,routine CV did not slow or impede biofilm development, as theanodic current after such analyses was 100.1% 3.4% (n � 6)of the current recorded before analyses. This was not unex-pected, as Geobacter spp. are commonly exposed to metals inthis potential range. Future work with stricter anaerobes, ororganisms adapted to higher-potential environments, shouldindependently verify the potential range which is not harmfulto electron transfer.

Stable catalytic features with a high signal-to-noise ratiowere observed when CV was performed at low scan rates (1mV/s). Under these conditions, the rate of electron transferrose rapidly at a potential higher than �0.3 V, reaching alimiting current above a potential of �0.1 V (Fig. 4). Thisresponse, commonly called a “reversible catalytic wave” (7),indicated continuous regeneration of the entire series of pro-teins catalyzing electron transfer from the substrate to theelectrode. In preliminary investigations, each increase in thescan rate significantly increased the ohmic current and shiftedthe location of peaks in voltammetric waves, as expected fromkinetic effects (see Fig. S2 in the supplemental material). How-ever, increasing the scan rate by fivefold did not significantlyalter the limiting current, demonstrating that at low scan rates,conditions were within a timescale sufficient to establish sus-tained catalysis by cells at each applied potential. Since themost informative scan rate could vary for each experimentalsetup, scan rates used in this report should be taken only as anaid in choosing conditions for new organism or electrode sur-face studies.

The inflection point(s) in such catalytic waves was visualizedas a peak using first-derivative analysis of voltammetry data(using software described in Materials and Methods) (2, 16).Such analysis of four independent biofilms is shown in Fig. 4.One dominant inflection point could be detected by this anal-ysis, centered at about �0.15 V, which increased in height withthe age of electrode-attached culture, while smaller intensitypeaks, centered at �0.220 V and �0.020 V versus SHE, couldalso be detected. The dominant peak revealed in derivativeanalyses was consistent with the presence of a single oxidation/reduction event in electron transfer that was rate limiting (1, 9,15). As shown in Fig. 4, these values were highly repeatablebetween individual biological replicates.

Once a biocatalyst is adsorbed, it is also possible to deter-mine its response to increasing concentrations of substrate(24). To test this approach, the electron donor was first re-

FIG. 3. SEMs of glassy carbon polished with P400 versus P4000 grittreatment (top). Bar graph (below) shows maximum current measured(n � 2 for each roughness) during CA after 72 h of growth. Alsoindicated (f, right y axis) is the average grit size of each polishingtreatment.

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moved from the medium, and electrodes were incubated at0.24 V for 36 h until no residual anodic (oxidation) current wasobserved. At this point, the effect of lowering substrate con-centrations could be determined. The anodic-limiting currentat �0.1 V was determined from CV and was found to increaselinearly with the acetate concentration when acetate was 200�M. At acetate concentrations higher than 3 mM, the anodic-limiting current did not change (Fig. 5). The midpoint poten-tial of catalytic waves remained centered at the same positionfor all concentrations, suggesting a similar electron transfermechanism and rate-controlling reaction, even at low currentdensities. This dose-response test detected acetate at concen-trations as low as 3.5 �M (signal-to-noise ratio, �3).

DPV. Although CV is the most informative method used toinvestigate catalytic substrate oxidation by adsorbed enzymes,it has a low detection limit, and subtraction of the ohmic(capacitive) current is necessary to reveal small features (9).Furthermore, when electron transfer between adsorbed en-zymes and electrodes is slow, as is expected for complex elec-tron transfer chains studied in stationary electrodes, CV re-

quires substantial time, and proper derivative analysis requirespostprocessing of data (40). In comparison, pulse methodshave the potential to reveal characteristic peaks while cancel-ling out capacitive current, even at higher scan rates, and areoften used as complementary techniques to CV (20, 44, 65).

Preliminary experiments with G. sulfurreducens biofilms in-dicated that DPV could also be used to monitor biofilms non-destructively, across a range of scan rates (up to 50 mV/s) andpulse heights (up to 100 mV). The parameters chosen (seeMaterials and Methods) represent a compromise between thetime of analysis and sensitivity. When performed on maturebiofilms, DPV always revealed a broad peak, which increasedin height with the age of the biofilm. These voltammogramswere highly reproducible, with the peak centered at �0.105 0.005 V versus SHE (Fig. 6).

Additional peaks at lower and higher potential with respectto the main peak were also detected. The results were consis-tent with multiple redox centers exposed on the surfaces of G.sulfurreducens cells that spanned a range of potentials (fromapproximately �0.2 to �0.1 V). In recent studies withShewanella biofilms grown on similar electrodes, we observedstrikingly different behavior, as DPV revealed a large, lessconvoluted peak at �0.21 V. These Shewanella cells werefound to secrete redox-active flavins that mediated electrontransfer (47), and changes in the DPV peak height could beused to monitor both the accumulation of this redox-activemediator at the electrode and its loss when the medium waschanged to remove soluble compounds. As the DPV peakheight was unaffected by medium changes in experiments withGeobacter biofilms, this represented additional support for ac-cumulation of an attached electron transfer agent and lack ofsoluble mediators within G. sulfurreducens biofilms. These re-sults demonstrate the utility of combining sweep and pulsemethods in the study of living systems.

EIS. While originally applied to investigation of solid-solidinterfaces, EIS is today the most common alternating currentmethod applied to the characterization of aqueous biointer-

FIG. 4. (A) Low scan rate CV of G. sulfurreducens biofilms afterinoculation (0 h) and at maximum current production (72 h). (B) Firstderivatives of cyclic voltammograms of biofilms (P400 grit polishingtreatment) (n � 4) showing the midpoint potential detectable in cat-alytic waves of mature biofilms.

FIG. 5. Low scan rate CV (1 mV/s) of G. sulfurreducens at differentconcentrations of electron donor. The biofilm was poised at oxidizingpotential without an electron donor until no anodic current was ob-served. Then electron donor was added, and the limiting current wasrecorded (inset). The midpoint potential of catalytic curves was con-stant with increasing concentration of the electron donor.

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faces (10, 13). In most EIS methods, a small sinusoidal poten-tial perturbation is applied to the sample. The frequency of thisperturbation is changed in the range between few mHz and 105

Hz. The resulting sinusoidal current is analyzed via fast Fouriertransform techniques to calculate the impedance (Z) of theinterface in the frequency domain to estimate charge transferresistance (32), diffusion at surfaces covered by protein mono-layers (20), charge transfer time constants (59), and mecha-nisms of electron transfer (8).

Applications of EIS are numerous in microbially influencedcorrosion, in which the sample is studied at the open circuitpotential (42, 51). However, we have shown that electrontransfer processes by G. sulfurreducens are voltage dependent(example shown in Fig. 3). Thus, while EIS is typically per-formed at open-circuit potentials, potentiostatic EIS (wherethe sinusoidal potential is applied while maintaining a givenpotential at the working electrode) was more likely to revealelectron transfer characteristics of active cells attached to elec-trodes.

Figure 7 shows a comparison of a glassy carbon electrode

before and after colonization by G. sulfurreducens. When EISwas performed at �0.16 V versus SHE, the presence of G.sulfurreducens reduced the charge transfer resistance (inferredfrom the Z value at low frequencies), with a main time con-stant in the 1 to 10 Hz frequency range. At high rates ofelectron transfer, a second, slower time constant was observedin the 0.01 to 0.1 Hz range. The addition of a second timeconstant to the model did not significantly improve the disper-sion of residuals, likely due to the low frequency of the secondtime constant.

The importance of potentiostatic EIS was illustrated by thesignificant influence that the imposed potential had on thecharge transfer resistance and time constants for G. sulfurre-ducens biofilms (Fig. 8; Table 1). When the imposed potentialwas similar to the midpoint potential of the catalytic waveobserved in CV, the lowest resistances and time constants weremeasured. At potentials outside of this range, differences incharge transfer were much smaller (in agreement with CVresults), resulting in higher apparent values for the chargetransfer resistance and time constant. Across the potentialrange explored, the solution resistance (R � 17.2 0.4 �),double-layer capacitance (Cp � 3.0 0.9 mF), and nonidealitycoefficient (a � 0.72 0.06) did not change appreciably. Theseresults again highlighted how data from one method (CV)could be used to select parameters in a complementary method(EIS). They also demonstrated the importance of investigatingthe proper analysis parameters for each bacterial system, asanother strain with a different midpoint potential for its cata-lytic process would need to be studied at a different imposedvoltage to obtain the fastest time constant.

Implications. Many studies providing voltammetry datafrom bacterium-electrode interactions have now been pub-lished. While reports differ in their pretreatment of bacteria,biofilm age, growth medium, environmental conditions, elec-trode materials, and electrochemical parameters, a generaltrend of catalytic wave-like behavior is clear (14, 31, 53, 68),showing that many bacteria and communities behave in a man-

FIG. 6. (A) DPV (scan rate, 4 mV/s; pulse height, 50 mV) undercatalytic (presence of acetate) conditions at inoculation and after 72 hof growth. (B) The shape of peaks and average potential for DPV ofmature biofilms (72 h) between four independent biological replicates.

FIG. 7. EIS of G. sulfurreducens biofilms at V � �0.16 V versusSHE; modulus of impedance (Z) after inoculation (F) and at 72 h (E);phase angle V-I after inoculation (f) and at 72 h (�). Biofilm devel-opment results in lower charge transfer resistance at the electrode-biofilm interface, with two distinct charge transfer processes around 1and 0.03 Hz.

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ner similar to the early stage Geobacter biofilms grown ongeometrically simple electrodes shown in this work. As a result,we can confirm what early observations suggested, thatGeobacter bacteria behave as attached biocatalysts, stimulatedby changes across a narrow potential range. For example, in aprior study, where we adsorbed washed Geobacter cells oncarbon paper electrodes (using pectin to retain free cells) (62),similar features were reported even after a few hours of cellsbeing exposed to electrodes. These consistencies further sug-gest that these responses and potentials are not dramaticallyinfluenced by such factors as stage of growth or electrodematerial.

In particular, this study shows how the response of an intactfilm to a range of applied potentials can be measured system-atically and analyzed to produce data that are easily comparedbetween species, conditions, or electrode surfaces. As scanrates are lowered, electron transfer kinetic effects (rates ofelectron transfer between cell surfaces and electrodes) areminimized, and enzymatic effects (related to microorganisms’

activity) become primary factors. As the goal was to initiallycharacterize the catalytic behavior of the system, low scan rates(1 mV/s) were chosen, since they permitted reactions with atime constant on the order of �1 s to be active as turnoverprocesses at each imposed potential step. At high levels ofelectron donor and low scan rates, catalytic voltammogramsshould therefore be representative of steady-state conditions.In addition, minimization of ohmic current aided in identifi-cation of inflection points in derivative analysis. The choice ofa relatively low scan rate was also supported by data from EISmeasurements, in which we found time constants on the orderof 0.1 and 1 to 10 s. For a system such as this, with whatappears to be sluggish interfacial electron transfer kinetics, thepotential difference between anodic and cathodic peaks for agiven redox couple could change significantly with even modestchanges in scan rate. Future work will focus on the use ofelectrochemical techniques under nonturnover conditions tobetter elucidate these electron transfer kinetics for a morecomplete understanding of the interplay between microbialcatalytic abilities and interfacial electron transfer.

The high reproducibility between biological replicates andthe ability to perform experiments with a range of electrodesprovided a robust measurement of G. sulfurreducens currentdensities, which approached 2 A/m2 for electrodes polishedwith 35-�m-grit-size particles. The catalytic wave consistentlyobserved for G. sulfurreducens is an independent demonstra-tion that interior oxidative processes of this organism arelinked via a continuous pathway to surfaces and that the entirecollection of attached organisms (i.e., the biofilm) behaves asan adsorbed catalyst. The midpoint potential of the catalyticwave at �0.15 V supports a model with a dominant rate-limiting electron transfer reaction and shows that G. sulfurre-ducens respiration rate does not increase when cells are pro-vided with an electron acceptor with a potential greater than 0V. The latter result implies that the final step of electrontransfer (e.g., between a terminal external protein and theelectrode) is not rate limiting, as this process can always beaccelerated by additional applied potential (as described by thestandard rate law kf � k0 e�E) (9). The midpoint and thelimiting current potential found in this study are consistentwith G. sulfurreducens being adapted for the reduction of ironoxides with a potential between �0.2 and 0 V versus SHE inthe environment and suggest that cells do not derive any ad-ditional energetic benefit from higher-potential electron ac-ceptors.

While the study of an intact electron transport chain in G.sulfurreducens represents perhaps a complex example of thiselectrochemical approach, similar methods have been used tocharacterize extracts from bacterial cells adsorbed onto, or

FIG. 8. EIS of mature G. sulfurreducens biofilms (72 h) performedat different potentials. (A) Bode plot. Modulus of impedance (Z) at0.04 (filled black circles), �0.06 (filled red circles), �0.16 (filled bluecircles), and �0.26 V (filled green circles) versus SHE; phase angle V-Iat 0.04 (open black circles), �0.06 (open red circles), �0.16 (open bluecircles), and �0.26 V (open green circles) versus SHE. Note thatfrequency of the major charge transfer process changes with appliedpotential. The second charge transfer process can be observed only at�0.16 V. (B) Nyquist plot. The relaxation times for the electron trans-fer process at the interface are summarized in Table 1.

TABLE 1. EIS parameters for biofilms at 72 ha

Potential vsSHE (V)

Charge transferresistance (�)

Characteristicfrequency (Hz)

Timeconstant (s)

�0.042 23,500 0.01 �100�0.058 1,050 0.032 32�0.158 204 0.5 2�0.258 2,440 0.04 25

a Data were fitted with the single time constant model described in Materialsand Methods. Values are averages of data from two separate experiments.

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incorporated into, electrode materials (26, 45, 46, 61). In ad-dition, researchers have washed and concentrated whole cellsfor brief voltammetry analyses and detected possible redox-active agents on the external surfaces of cells (39). The benefitsof using living cells in the work reported here include the lackof a protein purification step and the fact that cells are allowedto assemble the actual biofilm structure responsible for thisunique extracellular electron transfer process.

The multiple peaks visible in both CV and DPV analysesalso confirm the complexity of the G. sulfurreducens surface.Multiple cytochromes and redox proteins have been previouslyimplicated in outer membrane-based electron transfer in pro-teomic and labeling studies (23, 36, 41). As most proteinsimplicated in electron transport by G. sulfurreducens containmultiple hemes or redox centers, the detected redox centerscould reflect individual hemes, domains that act as a singlecenter, or individual proteins. Recent work with the multihemecytochrome MtrC (30) showed that multiheme proteins do notdemonstrate classic, individual redox behavior for each hemebut rather act as a cluster with a broader midpoint. In anotherstudy (67), the same protein was observed to behave as twopentaheme domains with broad midpoint potentials. Futurework with specific mutants lacking key redox proteins in G.sulfurreducens will aid in identifying the origin of these peaks.

Based on the results reported here, voltammetric methodspreviously developed to characterize electron transfer phe-nomena by enzymes adsorbed at carbon electrodes can beextended to the characterization of viable biofilms. By choos-ing the appropriate conditions, these methods are not destruc-tive and allow in vivo determination of electron transfer fromwhole cells to electrodes under conditions that are comparableto those encountered in natural environments. Both thermo-dynamic and kinetic parameters can be determined and usedto define the phenotype of an organism for comparison withother strains or mutants. The proposed methods can be ap-plied to well-defined pure cultures, as well as to complex mi-crobial communities, and could allow for quantitative compar-isons in the development of better microbial catalysts based ondirect electron transfer between bacteria and electrodes.

ACKNOWLEDGMENTS

We thank IREE and NSF (grant DBI-0454861) for their support, aswell as the BioTechnology Institute at the University of Minnesota.

We also thank Jeremy Alley for technical help with the experimentalsetup, Dirk Heering for suggestions in the interpretation of data, andLewis Werner for the SEM images of bare electrodes.

REFERENCES

1. Anderson, L. J., D. J. Richardson, and J. N. Butt. 2000. Using direct elec-trochemistry to probe rate limiting events during nitrate reductase turnover.Faraday Discuss. 116:155–169.

2. Angove, H. C., J. A. Cole, D. J. Richardson, and J. N. Butt. 2002. Protein filmvoltammetry reveals distinctive fingerprints of nitrite and hydroxylaminereduction by a cytochrome c nitrite reductase. J. Biol. Chem. 277:23374–23381.

3. Armstrong, F. A. 2002. Insights from protein film voltammetry into mecha-nisms of complex biological electron-transfer reactions. J. Chem. Soc. DaltonTrans. 5:661–671.

4. Armstrong, F. A. 2002. Protein film voltammetry: revealing the mechanismsof biological oxidation and reduction. Russ. J. Electrochem. 38:49–62.

5. Armstrong, F. A. 2005. Recent developments in dynamic electrochemicalstudies of adsorbed enzymes and their active sites. Curr. Opin. Chem. Biol.9:110–117.

6. Armstrong, F. A., A. M. Bond, F. N. Buchi, A. Hamnett, H. A. O. Hill, A. M.Lannon, O. C. Lettington, and C. G. Zoski. 1993. Electrocatalytic reduction

of hydrogen-peroxide at a stationary pyrolytic-graphite electrode surface inthe presence of cytochrome-C peroxidase—a description based on a micro-electrode array model for adsorbed enzyme molecules. Analyst 118:973–978.

7. Armstrong, F. A., H. A. Heering, and J. Hirst. 1997. Reactions of complexmetalloproteins studied by protein-film voltammetry. Chem. Soc. Rev. 26:169–179.

8. Bai, X., S. C. Dexter, and G. W. Luther III. 2006. Application of EIS withAu-Hg microelectrode in determining electron transfer mechanism. Electro-chim. Acta 51:1524–1533.

9. Bard, A. J., and L. R. Faulkner. 2001. Electrochemical methods. Wiley, NewYork, NY.

10. Barsoukov, E., and J. R. Macdonald. 2005. Impedance spectroscopy—the-ory, experiment, and applications. Wiley Interscience, Hoboken, NJ.

11. Bond, D. R., and D. R. Lovley. 2003. Electricity production by Geobactersulfurreducens attached to electrodes. Appl. Environ. Microbiol. 69:1548–1555.

12. Bond, D. R., and D. R. Lovley. 2005. Evidence for involvement of an electronshuttle in electricity generation by Geothrix fermentans. Appl. Environ.Microbiol. 71:2186–2189.

13. Brabec, V., V. Vetterl, and O. Vrana. 1996. Electroanalysis of biomacromol-ecules. Bioelectrochemistry: principles and practice, p. 287–359. In V. Bra-bec, D. Walz, and G. Milazzo (ed.), Experimental techniques in bioelectro-chemistry, vol. 3. Birkhauser Verlag, Basel, Switzerland.

14. Busalmen, J. P., A. Berna, and J. M. Feliu. 2007. Spectroelectrochemicalexamination of the interaction between bacterial cells and gold electrodes.Langmuir 23:6459–6466.

15. Butt, J. N., and F. A. Armstrong. 2008. Voltammetry of adsorbed redoxenzymes: mechanisms in the potential dimension, p. 91–128. In O. Hammerichand J. Ulstrup (ed.), Bioinorganic electrochemistry. Springer, Dordrecht, TheNetherlands.

16. Butt, J. N., J. Thornton, D. J. Richardson, and P. S. Dobbin. 2000. Volta-mmetry of a flavocytochrome c3: the lowest potential heme modulates fu-marate reduction rates. Biophys. J. 78:1001–1009.

17. Calvo, E. J., F. Battaglini, C. Danilowicz, A. Wolosiuk, and M. Otero. 2000.Layer-by-layer electrostatic deposition of biomolecules on surfaces for mo-lecular recognition, redox mediation and signal generation. Faraday Discuss.116:47–65.

18. Chang, I. S., H. Moon, O. Bretschger, J. K. Jang, H. I. Park, K. H. Nealson,and B. H. Kim. 2006. Electrochemically active bacteria (EAB) and mediator-less microbial fuel cells. J. Microbiol. Biotechnol. 16:163–177.

19. Chaudhuri, S. K., and D. R. Lovley. 2003. Electricity generation by directoxidation of glucose in mediator-less microbial fuel cells. Nat. Biotechnol.21:1229–1232.

20. Chi, Q., J. Zhang, J. U. Nielsen, E. P. Friis, I. Chorkendorff, G. W. Canters,J. E. T. Andersen, and J. Ulstrup. 2000. Molecular monolayers and interfa-cial electron transfer of Pseudomonas aeruginosa azurin on Au(111). J. Am.Chem. Soc. 122:4047–4055.

21. Chin, K. J., A. Esteve-Nunez, C. Leang, and D. R. Lovley. 2004. Directcorrelation between rates of anaerobic respiration and levels of mRNA forkey respiratory genes in Geobacter sulfurreducens. Appl. Environ. Microbiol.70:5183–5189.

22. de Beer, D., P. Stoodley, F. Roe, and Z. Lewandowski. 1994. Effects of biofilmstructures on oxygen distribution and mass transport. Biotechnol. Bioeng.43:1131–1138.

23. Ding, Y. H. R., K. K. Hixson, C. S. Giometti, A. Stanley, A. Esteve-Nunez, T.Khare, S. L. Tollaksen, W. H. Zhu, J. N. Adkins, M. S. Lipton, R. D. Smith,T. Mester, and D. R. Lovley. 2006. The proteome of dissimilatory metal-reducing microorganism Geobacter sulfurreducens under various growth con-ditions. Biochim. Biophys. Acta 1764:1198–1206.

24. Elliott, S. J., K. R. Hoke, K. Heffron, M. Palak, R. A. Rothery, J. H. Weiner,and F. A. Armstrong. 2004. Voltammetric studies of the catalytic mechanismof the respiratory nitrate reductase from Escherichia coli: how nitrate reduc-tion and inhibition depend on the oxidation state of the active site. Biochem-istry 43:799–807.

25. Fan, Y. Z., H. Hu, and H. Liu. 2007. Enhanced Coulombic efficiency andpower density of air-cathode microbial fuel cells with an improved cellconfiguration. J. Power Sources 171:348–354.

26. Garjonyte, R., and A. Malinauskas. 2003. Investigation of baker’s yeastSaccharomyces cerevisiae- and mediator-based carbon paste electrodes asamperometric biosensors for lactic acid. Sens. Actuators B 96:509–515.

27. Giao, M. S., M. I. Montenegro, and M. J. Vieira. 2003. Monitoring biofilmformation by using cyclic voltammetry—effect of the experimental conditionson biofilm removal and activity. Water Sci. Technol. 47(5):51–56.

28. Giao, M. S., M. I. Montenegro, and M. J. Vieira. 2005. The influence ofhydrogen bubble formation on the removal of Pseudomonas fluorescens bio-films from platinum electrode surfaces. Process Biochem. 40:1815–1821.

29. Gil, G. C., I. S. Chang, B. H. Kim, M. Kim, J. K. Jang, H. S. Park, and H. J.Kim. 2003. Operational parameters affecting the performance of a mediator-less microbial fuel cell. Biosens. Bioelectron. 18:327–334.

30. Hartshorne, R. S., B. N. Jepson, T. A. Clarke, S. J. Field, J. Fredrickson, J.Zachara, L. Shi, J. N. Butt, and D. J. Richardson. 2007. Characterization ofShewanella oneidensis MtrC: a cell-surface decaheme cytochrome involved in

7336 MARSILI ET AL. APPL. ENVIRON. MICROBIOL.

on January 12, 2021 by guesthttp://aem

.asm.org/

Dow

nloaded from

respiratory electron transport to extracellular electron acceptors. J. Biol.Inorg. Chem. 12:1083–1094.

31. He, Z., S. D. Minteer, and L. T. Angenent. 2005. Electricity generation fromartificial wastewater using an upflow microbial fuel cell. Environ. Sci. Tech-nol. 39:5262–5267.

32. He, Z., N. Wagner, S. D. Minteer, and L. T. Angenent. 2006. An upflowmicrobial fuel cell with an interior cathode: assessment of the internal re-sistance by impedance spectroscopy. Environ. Sci. Technol. 40:5212–5217.

33. Heering, H. A., J. Hirst, and F. A. Armstrong. 1998. Interpreting the catalyticvoltammetry of electroactive enzymes adsorbed on electrodes. J. Phys.Chem. B 102:6889–6902.

34. Hernandez, M. E., A. Kappler, and D. K. Newman. 2004. Phenazines andother redox-active antibiotics promote microbial mineral reduction. Appl.Environ. Microbiol. 70:921–928.

35. Holmes, D. E., J. S. Nicoll, D. R. Bond, and D. R. Lovley. 2004. Potential roleof a novel psychrotolerant member of the family Geobacteraceae, Geopsy-chrobacter electrodiphilus gen. nov., sp nov. in electricity production by amarine sediment fuel cell. Appl. Environ. Microbiol. 70:6023–6030.

36. Holmes, D. E., S. K. Chaudhuri, K. P. Nevin, T. Mehta, B. A. Methe, A. Liu,J. E. Ward, T. L. Woodard, J. Webster, and D. R. Lovley. 2006. Microarrayand genetic analysis of electron transfer to electrodes in Geobacter sulfurre-ducens. Environ. Microbiol. 8:1805–1815.

37. Jang, J. K., T. H. Pham, I. S. Chang, K. H. Kang, H. Moon, K. S. Cho, andB. H. Kim. 2004. Construction and operation of a novel mediator- andmembrane-less microbial fuel cell. Process Biochem. 39:1007–1012.

38. Kim, B. C., C. Leang, Y. H. R. Ding, R. H. Glaven, M. V. Coppi, and D. R.Lovley. 2005. OmcF, a putative c-type monoheme outer membrane cyto-chrome required for the expression of other outer membrane cytochromes inGeobacter sulfurreducens. J. Bacteriol. 187:4505–4513.

39. Kim, H. J., H. S. Park, M. S. Hyun, I. S. Chang, M. Kim, and B. H. Kim.2002. A mediator-less microbial fuel cell using a metal reducing bacterium,Shewanella putrefaciense. Enzyme Microb. Technol. 30:145–152.

40. Koide, S., and K. Yokoyama. 1999. Electrochemical characterization of anenzyme electrode based on a ferrocene-containing redox polymer. J. Elec-troanal. Chem. 468:193–201.

41. Leang, C., L. A. Adams, K. J. Chin, K. P. Nevin, B. A. Methe, J. Webster,M. L. Sharma, and D. R. Lovley. 2005. Adaptation to disruption of theelectron transfer pathway for Fe(III) reduction in Geobacter sulfurreducens.J. Bacteriol. 187:5918–5926.

42. Lee, A. K., M. G. Buehler, and D. K. Newman. 2006. Influence of a dual-species biofilm on the corrosion of mild steel. Corrosion Sci. 48:165–178.

43. Liu, H., R. Ramnarayanan, and B. E. Logan. 2004. Production of electricityduring wastewater treatment using a single chamber microbial fuel cell.Environ. Sci. Technol. 38:2281–2285.

44. Liu, H. H., J. L. Lu, M. Zhang, D. W. Pang, and H. D. Abruna. 2003. Directelectrochemistry of cytochrome c surface confined on DNA-modified goldelectrodes. J. Electroanal. Chem. 544:93–100.

45. Lojou, E., M. C. Durand, A. Dolla, and P. Bianco. 2002. Hydrogenase activitycontrol at Desulfovibrio vulgaris cell-coated carbon electrodes: biochemicaland chemical factors influencing the mediated bioelectrocatalysis. Electro-analysis 14:913–922.

46. Lojou, E., M. T. Gludici-Orticoni, and P. Bianco. 2005. Direct electrochem-istry and enzymatic activity of bacterial polyhemic cytochrome c3 incorpo-rated in clay films. J. Electroanal. Chem. 579:199–213.

47. Marsili, E., D. B. Baron, I. Shikhare, D. Coursolle, J. Gralnick, and D. R.Bond. 2008. Shewanella secretes flavins that mediate extracellular electrontransfer. Proc. Natl. Acad. Sci. USA 105:3968–3973.

48. Mehta, T., M. V. Coppi, S. E. Childers, and D. R. Lovley. 2005. Outermembrane c-type cytochromes required for Fe(III) and Mn(IV) oxide re-duction in Geobacter sulfurreducens. Appl. Environ. Microbiol. 71:8634–8641.

49. Minteer, S. D., B. Y. Liaw, and M. J. Cooney. 2007. Enzyme-based biofuelcells. Curr. Opin. Biotechnol. 18:228–234.

50. Park, D. H., and J. G. Zeikus. 2000. Electricity generation in microbial fuelcells using neutral red as an electronophore. Appl. Environ. Microbiol.66:1292–1297.

51. Perez, E. J., R. Cabrera-Sierra, I. Gonzalez, and F. Ramirez-Vives. 2007.Influence of Desulfovibrio sp. biofilm on SAE 1018 carbon steel corrosion insynthetic marine medium. Corrosion Sci. 49:3580–3597.

52. Perez-Roa, R. E., D. T. Tompkins, M. Paulose, C. A. Grimes, M. A. Ander-son, and D. R. Noguera. 2006. Effects of localised, low-voltage pulsed electricfields on the development and inhibition of Pseudomonas aeruginosa bio-films. Biofouling 22:383–390.

53. Prasad, D., S. Arun, A. Murugesan, S. Padmanaban, R. S. Satyanarayanan,S. Berchmans, and V. Yegnaraman. 2007. Direct electron transfer with yeastcells and construction of a mediatorless microbial fuel cell. Biosens. Bio-electron. 22:2604–2610.

54. Qian, X. L., G. Reguera, T. Mester, and D. R. Lovley. 2007. Evidence thatOmcB and OmpB of Geobacter sulfurreducens are outer membrane surfaceproteins. FEMS Microbiol. Lett. 277:21–27.

55. Rabaey, K., N. Boon, M. Hofte, and W. Verstraete. 2005. Microbial phen-azine production enhances electron transfer in biofuel cells. Environ. Sci.Technol. 39:3401–3408.

56. Rabaey, K., N. Boon, S. D. Siciliano, M. Verhaege, and W. Verstraete. 2004.Biofuel cells select for microbial consortia that self-mediate electron trans-fer. Appl. Environ. Microbiol. 70:5373–5382.

57. Reguera, G., K. D. McCarthy, T. Mehta, J. S. Nicoll, M. T. Tuominen, andD. R. Lovley. 2005. Extracellular electron transfer via microbial nanowires.Nature 435:1098–1101.

58. Reguera, G., K. P. Nevin, J. S. Nicoll, S. F. Covalla, T. L. Woodard, and D. R.Lovley. 2006. Biofilm and nanowire production leads to increased current inGeobacter sulfurreducens fuel cells. Appl. Environ. Microbiol. 72:7345–7348.

59. Repo, T., G. Zhang, A. Ryyppo, and R. Rikala. 2000. The electrical imped-ance spectroscopy of Scots pine (Pinus sylvestris L.) shoots in relation to coldacclimation. J. Exp. Bot. 51:2095–2107.

60. Rhoads, A., H. Beyenal, and Z. Lewandowski. 2005. Microbial fuel cell usinganaerobic respiration as an anodic reaction and biomineralized manganeseas a cathodic reactant. Environ. Sci. Technol. 39:4666–4671.

61. Sallez, Y., P. Bianco, and E. Lojou. 2000. Electrochemical behavior of c-typecytochromes at clay-modified carbon electrodes: a model for the interactionbetween proteins and soils. J. Electroanal. Chem. 493:37–49.

62. Srikanth, S., E. Marsili, M. C. Flickinger, and D. R. Bond. 2008. Electro-chemical characterization of Geobacter sulfurreducens cells immobilized ongraphite paper electrodes. Biotechnol. Bioeng. 99:1065–1073.

63. Taillefert, M., J. S. Beckler, E. Carey, J. L. Burns, C. M. Fennessey, and T. J.DiChristina. 2007. Shewanella putrefaciens produces an Fe(III)-solubilizingorganic ligand during anaerobic respiration on insoluble Fe(III) oxides.J. Inorg. Biochem. 101:1760–1767.

64. von Canstein, H., J. Ogawa, S. Shimizu, and J. R. Lloyd. 2008. Secretion offlavins by Shewanella species and their role in extracellular electron transfer.Appl. Environ. Microbiol. 74:615–623.

65. Wang, S. G., R. L. Wang, P. J. Sellin, and Q. Zhang. 2004. DNA biosensorsbased on self-assembled carbon nanotubes. Biochem. Biophys. Res. Com-mun. 325:1433–1437.

66. White, D. 2000. The physiology and chemistry of prokaryotes. Oxford Uni-versity Press, New York, NY.

67. Wigginton, N. S., K. M. Rosso, and M. F. Hochella. 2007. Mechanisms ofelectron transfer in two decaheme cytochromes from a metal-reducing bac-terium. J. Phys. Chem. B 111:12857–12864.

68. Yoon, S. M., C. H. Choi, M. Kim, M. S. Hyun, S. H. Shin, D. H. Yi, and H. J.Kim. 2007. Enrichment of electrochemically active bacteria using a three-electrode electrochemical cell. J. Microbiol. Biotechnol. 17:110–115.

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