1 1 2 3 4 Microbial biofilm voltammetry: direct electrochemical 5 ...

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Microbial biofilm voltammetry: direct electrochemical 5

characterization of catalytic electrode-attached biofilms 6

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Enrico Marsili1, Janet B. Rollefson2, Daniel B. Baron1, Raymond M. Hozalski3, and 10

Daniel R. Bond1,4* 11

12 1BioTechnology Institute, University of Minnesota, St. Paul, Minnesota 55108 13 2Department of Biochemistry, Molecular Biology and Biophysics, University of 14

Minnesota, Minneapolis, Minnesota 55455 15 3Department of Civil Engineering, University of Minnesota, Minneapolis, Minnesota 16

55455 17 4 Department of Microbiology, University of Minnesota, Minneapolis, Minnesota 55455 18

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Running title: Microbial biofilm voltammetry 26

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Keywords: Geobacter, voltammetry, biofilms 28

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*Corresponding author. Mailing address: 39

BioTechnology Institute, University of Minnesota 40

140 Gortner Laboratory, 41

1479 Gortner Ave, 42

St. Paul, MN 55108 43

Phone: 612-624-8619, Fax: 612-625-1700. 44

Email: [email protected] 45

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Copyright © 2008, American Society for Microbiology and/or the Listed Authors/Institutions. All Rights Reserved.Appl. Environ. Microbiol. doi:10.1128/AEM.00177-08 AEM Accepts, published online ahead of print on 10 October 2008

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ABSTRACT 1

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While electrochemical characterization of enzymes immobilized on electrodes 3

has become common, there is still a need for reliable quantitative methods for study of 4

electron transfer between living cells and conductive surfaces. This work describes 5

growth of thin (<20 µm) Geobacter sulfurreducens biofilms on polished glassy carbon 6

electrodes, using stirred 3-electrode anaerobic bioreactors controlled by potentiostats 7

and non-destructive voltammetry techniques for characterization of viable biofilms. 8

Routine in vivo analysis of electron transfer between bacterial cells and electrodes was 9

performed, providing insight into the main redox-active species participating in electron 10

transfer to electrodes. At slow scan rates, cyclic voltammetry (CV) revealed catalytic 11

electron transfer between cells and the electrode, similar to what has been observed for 12

pure enzymes attached to electrodes under continuous turnover conditions. Differential 13

pulse voltammetry (DPV) and electrochemical impedance spectroscopy (EIS) also 14

revealed features that were consistent with electron transfer being mediated by an 15

adsorbed catalyst. Multiple redox-active species were detected, revealing complexity at 16

the outer surface of this bacterium. These techniques provide the basis for cataloging 17

quantifiable, defined electron transfer phenotypes as a function of potential, electrode 18

material, growth phase, and culture conditions, and provide a framework for 19

comparisons with other species or communities. 20

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INTRODUCTION 1

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Anaerobic metal-reducing Geobacteraceae catalyze the transfer of electrons 3

from reduced electron donors, such as acetate, to oxidized electron acceptors such as 4

Fe(III). While electron transport is confined to the inner membrane of most respiratory 5

bacteria (66), Fe(III) and Mn(IV) primarily exist in the environment as insoluble 6

precipitates at circumneutral pH. To reduce minerals of varying crystallinity, redox 7

potential, and surface charge, metal-reducing bacteria must possess a complete 8

mechanism for transporting electrons across the inner membrane, periplasm, outer 9

membrane, and outer surface (23,38,48,54,57,58). Soluble redox-active shuttles or 10

chelators can facilitate respiration beyond the outer membrane in some organisms 11

(12,34,55,56,63), but no evidence for the secretion of such a compound has been 12

observed in Geobacteraceae. 13

This electron transfer capability has recently been utilized in devices known as 14

‘microbial fuel cells’ (MFCs), where electrodes serve as electron acceptors to support 15

growth (11,18,35,36,62). However, variations in electrode surfaces, operating 16

temperatures, electron donor concentrations and culture conditions limit comparison 17

between electrode-reducing bacteria. In addition, MFCs aim to maximize electrical 18

power production per unit volume, using large anodes consisting of carbon felt 19

(19,39,50) or carbon paper (43) in small reactors (with surface:volume ratios between 20

25 and 640 (19,25,29,43). In such systems the actual surface accessible to bacteria is 21

not known, multiple ‘dead zones’ or diffusion-limited regions exist, (19), high-surface 22

area electrodes increase capacitive current and reduce the sensitivity of analyses 23

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(53,68), and irregular surfaces prevent mathematical modeling or imaging of attached 1

biomass (22). 2

Electrochemical techniques typically used to study purified redox proteins (3,4,7) 3

and cell extracts (26,45) could likely be adapted for the study of metal-reducing 4

bacteria. While protein voltammetry is limited by the success of purification and 5

immobilization of enzymes (17,49), metal-reducing bacteria represent a self-assembling 6

multienzyme system that naturally interfaces with electrodes. In this report, we describe 7

procedures for analysis of G. sulfurreducens biofilms on small carbon electrodes of 8

defined roughness in potentiostat-controlled 3-electrode cells. Several direct current 9

(DC) and alternating current (AC) techniques could be applied to the characterization of 10

the biofilm-electrode interface, and produced insights into electron transfer mechanisms 11

between living cells and electrodes. Application of these techniques in routine 12

characterization of other living bacteria and biofilm communities will allow quantitative 13

comparisons between electron transfer rates and mechanisms used by these 14

organisms. 15

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MATERIALS AND METHODS 17

Bacterial strain and culture media - G. sulfurreducens strain PCA (ATCC #51573) 18

was sub-cultured in our laboratory at 30°C using a vitamin-free anaerobic medium 19

containing per liter: 0.38 g KCl, 0.2 g NH4Cl, 0.069 g NaH2PO4•H2O, 0.04 g 20

CaCl2•2H2O, 0.2 g MgSO4•7H2O; 10 ml of a mineral mix (containing per 1 l : 0.1 g 21

MnCl2•4H2O, 0.3 g FeSO4•7H2O, 0.17 g CoCl2•6H2O, 0.1 g ZnCl2, 0.04 g CuSO4•5H2O, 22

0.005 g AlK(SO4)2•12H2O, 0.005 g H3BO3, 0.09 g Na2MoO4, 0.12 g NiCl2, 0.02 g 23

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NaWO4•2H2O and 0.10 g Na2SeO4). Acetate was provided as an electron donor at 20 1

mM. All media were adjusted to pH 6.8 prior to addition of 2 g/l NaHCO3, and were 2

flushed with oxygen-free N2/CO2 (80/20 v/v) prior to sealing with butyl rubber stoppers 3

and autoclaving. All experiments were initiated by inoculating with 40% v/v of cells 4

within 3 h of reaching maximum optical density (OD>0.6) in the medium described. 5

Electrode preparation - Glassy carbon (Toko America, New York NY) was 6

machine-cut into 2 x 0.5 x 0.1 cm electrodes. Freshly cut glassy carbon electrodes 7

were polished using aluminum oxide-silicon carbide sandpaper with grit designation 8

between P400 and P4000 (3M, Minneapolis MN), as indicated in ISO/FEPA standard 9

(http://www.fepa-abrasives.com). Mirror polished electrode surfaces were obtained with 10

0.05 µm Alumina powder (CH Instruments, Austin TX). Polished electrodes were 11

sonicated to remove debris, soaked overnight in 1 N HCl to remove metals and other 12

contaminants, washed twice with acetone and deionized water to remove organic 13

substances, and stored in deionized water. After each experiment, electrode surfaces 14

were cleaned with an additional 1 N NaOH treatment (to remove biomass), and the 15

entire surface was refreshed through sandpaper polishing and cleaning as described, to 16

remove immobilized electron transfer agents. These working electrodes were attached 17

to 0.1 mm Pt wires via miniature nylon screws that ensured electrical contact throughout 18

the experiment. 19

Electrode cell assembly - Platinum wires from the working electrode were 20

inserted into heat-pulled 3 mm glass capillary tubes (Kimble, Vineland NJ) and soldered 21

inside the capillary to copper wires. Counter electrodes consisted of a 0.1 mm diameter 22

Pt wire (Sigma-Aldrich, St. Louis MO) that was also inserted into a 3 mm glass capillary 23

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and soldered to a copper wire. The resistance of each electrode assembly was 1

measured, and electrodes with a total resistance higher than 0.5 Ω were discarded. 2

Reference electrodes were connected to bioreactors via a salt bridge assembled from a 3

3 mm glass capillary and a 3 mm vycor frit (BioAnalytical Systems, West Lafayette IN). 4

Electrode capillaries were inserted through ports in a custom-made Teflon lid that was 5

sealed with an O-ring gasket. This lid fit onto a 20 ml conical electrochemical cell 6

(BioAnalytical Systems, West Lafayette IN), which had been previously washed in 3 N 7

HNO3 (Figure 1A). 8

After the addition of a small magnetic stir bar, the cell was autoclaved for 15 9

minutes. Following autoclaving, the salt bridge was filled with 0.1 M Na2SO4 in 1% agar. 10

A saturated calomel reference electrode (Fisher Scientific, Pittsburg PA) was placed at 11

the top of this agar layer, and covered in additional Na2SO4 to ensure electrical contact 12

(60). The reactors were placed in an in-house built water bath to maintain cells at 30°C 13

(which introduced less electrical noise compared to large circulators, incubators or 14

temperature-controlled rooms) (Figure 1B). To maintain the strict anaerobic conditions 15

required by bacteria, all reactors were operated under a constant flow of sterile 16

humidified N2:CO2 (80:20 v/v), which had been passed over a heated copper column to 17

remove trace oxygen. Each reactor was located above an independent magnetic 18

stirring unit. 19

Autoclaved bioreactors flushed free of oxygen, filled with sterile growth medium, 20

and incubated at 30°C were analyzed before each experiment, to verify anaerobicity 21

and the absence of redox active species. Electrochemical cells showing residual peaks 22

in differential pulse voltammetry (DPV), anodic current in cyclic voltammetry (CV), or 23

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baseline noise were discarded as having possible electrode cleanliness or connection 1

noise issues. These autoclaved, verified bioreactors were then used for growth of G. 2

sulfurreducens cultures. 3

A typical bioreactor was inoculated with 40% v/v of stationary phase G. 4

sulfurreducens cells. After inoculation, a potential step of 0.24 V vs. SHE was applied 5

and the reactors were incubated until current stabilized (~72 hours). The potential step 6

was ~200 mV higher than the midpoint potential sustaining a half-maximal anodic 7

current (as determined by preliminary CV, see below) (9). 8

Electrochemical instrumentation - A 16-channel potentiostat (VMP®, Bio-Logic 9

USA, Knoxville TN) was connected to the 3-electrode cells described above (Figure 10

1B). Software from the same producer (EC-Lab, V.9.41) was used to run simultaneous 11

multitechnique electrochemistry routines, which include CV, DPV, and 12

chronoamperometry (CA). Scan rates higher than 100 mV/s and all electrochemical 13

impedance spectroscopy (EIS) experiments were performed using a Gamry PCI4 14

Femtostat (Gamry Instruments, Warminster PA) and EIS data fitting was performed with 15

E-Chem Analyst, v. 5.1. Post-acquisition analysis of CV data was performed with the 16

software Utilities for Data Analysis (UTILS), kindly provided by Dr. D. Heering (version 17

1.0; University of Delft, The Netherlands [http://www.tudelft.nl/]). All measurements, 18

with exception of CA, were performed in succession without stirring enabled. The 19

parameters for each method were: DPV: Ei= -0.558 V vs. standard hydrogen electrode 20

(SHE), Ef= 0.242 V vs. SHE; pulse height 50 mV, pulse width 300 ms, step height 2 mV, 21

step time 500 ms; scan rate 4 mV/s, current averaged over the last 80% of the step (1 22

sec, 12 points), accumulation time 5 s. CV: equilibrium time 5 sec, scan rate 1 mV/s, Ei= 23

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-0.558 V vs. SHE, Ef= 0.242 V vs. SHE, current averaged over the whole step (1 sec, 1

10 points). CA: E = 0.242 V vs. SHE. 2

EIS was conducted while maintaining a DC voltage between the working 3

electrode and the reference electrode (potentiostatic EIS), as electron transfer 4

processes by Geobacter were observed to be voltage-dependent (62). EIS was 5

performed at four potentials: 0.04, -0.06, -0.16, -0.26 V vs. SHE, with a perturbation 6

amplitude of 10 mV. The frequency was varied from 105 to 0.01 Hz, as most biological 7

charge transfer phenomena are observed in this range (10). Data were then fitted with a 8

simple empirical model (62), using a constant phase element to account for irregularities 9

in the morphology and chemistry of the glassy carbon surface. EIS was typically only 10

performed at the end of incubations as extensive characterization required electrodes to 11

be held at multiple potentials for long periods of time. 12

Confocal and SEM analysis - A S3500N scanning electron microscope (SEM) 13

(Hitachi, Japan) was used to image freshly polished (SEM) electrodes. After being 14

thoroughly cleaned with deionized water, 2 x 0.5 x 0.1 cm graphite electrodes freshly 15

polished with P400 and P4000 sandpapers were air dried and fixed through graphite 16

tape to a sample stub. The sample stubs were placed directly into the SEM vacuum 17

compartment and examined at accelerating voltage of 5 kV. 18

A Nikon C1 Spectral Imaging Confocal Microscope (Nikon, Japan) with a 60x 19

lens was used to image biofilm-covered electrodes. Immediately following harvesting, 20

the biofilm-covered electrodes were gently washed in growth medium to remove 21

planktonic cells, then stained with the LIVE/DEAD Baclight Bacterial Viability Kit 22

(Invitrogen Corp., Carlsbad, CA), incubated for 15 minutes in the dark, rinsed with 23

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growth medium, and placed on a microscope slide. The coverslip was supported by 1

glass spacers thicker than the electrode, to prevent damage to biofilm structure. Two 2

laser wavelengths, 488 and 561 nm, were used to excite the two stains. 3

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RESULTS AND DISCUSSION 5

Colonization of electrodes. In microbial fuel cells, the half-cell potential at the 6

anode varies with the activity of the bacterial biofilm and the chosen external resistance 7

(37,43). Potentiostat controlled electrodes at a sufficiently positive potential are 8

equivalent to anodes in which electron acceptor is non-limiting, reducing the technical 9

complexity and simplifying the conceptual model of electron transfer. However, at high 10

oxidative potentials, conformational change, unfolding, and irreversible process could 11

alter the catalytic ability of enzymes, decreasing anodic (oxidation) current (27,52). In all 12

experiments described, the potential step used was 0.24 V vs. SHE. This potential was 13

chosen based on preliminary CV experiments, which showed that oxidation current from 14

cells was independent of applied potential in the range 0.1 – 0.3 V vs. SHE. 15

After inoculation, the potential step was applied, and a rapidly increasing anodic 16

(oxidation) current was detected (Figure 2). As this rate of increase was many times 17

faster than what could be explained by known growth rates of G. sulfurreducens, it was 18

likely due to cell attachment to the electrode. The attachment phase was most rapid 19

when fumarate-limited cells (compared to mid-log phase cells) were used as the 20

inoculum, consistent with reports linking acceptor limitation to the expression of 21

enzymes important to metal reduction (21,23). This was followed by a growth phase, 22

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characterized by an exponential increase in current, which doubled at a rate typically 1

observed for G. sulfurreducens reducing Fe(III)-citrate or fumarate (doubling time ~ 7 h). 2

Within 72 h, electrodes reached a saturation phase characterized by a stable 3

maximum electron transfer rate with a standard deviation between biological replicates 4

of less than 3%. Confocal imaging of electrodes always revealed uniform thin films of 5

~15 µm with few structural features such as pillars (Supplementary Figure 1). During the 6

attachment and growth phases, electrodes were tested for the effect of voltammetry 7

analyses on the rate of growth or current plateau. Repeated CV and DPV (see 8

methods), did not alter key midpoint potentials, colonization rates, or final current 9

densities (Figure 2). 10

In order to minimize contribution of other electron transfer agents, we used a 11

medium lacking redox-active species, such as vitamins (i.e., riboflavin), redox indicators 12

(resazurin), and chemical reductants (cysteine, DTT) (47,64). In our recent study (47), 13

we showed that flavin concentrations in the nM range can be detected under the 14

experimental conditions applied in the present work. When the medium in reactors was 15

replaced, removing planktonic cells and any potential microbially-produced soluble 16

electron transfer agent, the electron transfer rate to electrodes remained unchanged. In 17

addition, removal of NTA-chelated minerals from the medium (once electrode 18

colonization had occurred) did not affect current production. These findings created 19

stringent conditions for collection of baseline data in the absence of most common 20

soluble redox-active compounds. A systematic examination of the effects of commonly 21

present redox species on electron transfer by Geobacter will be the focus of a future 22

report. 23

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The effect of microscale surface roughness on bacterial attachment and direct 1

electron transfer has not been reported. Figure 3 shows the surface features of carbon 2

electrodes polished with different sandpapers. When G. sulfurreducens biofilms were 3

grown on electrodes polished with increasing grit size, the maximum current density 4

increased over 100% for the same geometric area (Figure 3). Polishing with particles 5

smaller than ~5 µm did not significantly reduce the maximum current. Therefore, even 6

µm-scale features have to be controlled to allow reproducibility and comparisons 7

between cultures and electrodes in this and future studies. 8

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Cyclic voltammetry (CV). CV is the principal diagnostic method in protein film 10

voltammetry, which studies immobilized enzymes on rotating and stationary carbon 11

electrodes. When in presence of a suitable substrate, and slowly cycled through a 12

range of applied potentials at low scan rates, enzymes have time to undergo multiple 13

turnovers at each sampled potential (5-7,33). As scan rates increase, slow reactions 14

may not have time to occur before the potential changes to the next step, and 15

concentration gradients representative of steady-state conditions may not have time to 16

establish. Thus, as scan rate increases, the kinetics of interfacial electron transfer 17

between redox protein(s) and electrodes affects more strongly the voltammetric 18

response, and can hide the kinetics of continuous enzymatic turnover. As the purpose 19

of this study was to establish methods for measuring catalytic (turnover) kinetics in 20

viable bacterial biofilms, in the absence of prior knowledge regarding interfacial electron 21

transfer rates, slow scan rates represented the most conservative experimental 22

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conditions. Parameters for the study of non-turnover voltammetry at high scan rates will 1

be discussed in subsequent reports. 2

Application of CV to living bacteria required limiting the potential range to prevent 3

harmful oxidizing or reducing conditions, as well as selection of informative scan rates. 4

While it is known that application of potentials can disrupt biofilms either by producing 5

hydrogen (28) or by inducing unfolding/oxidation of adsorbed proteins at oxidizing 6

potentials (52), G. sulfurreducens biofilms were not harmed by exposure to potentials 7

between -0.55 and 0.24 V vs. SHE. For example, routine CV did not slow or impede 8

biofilm development, as the anodic current after such analyses was 100.1%±3.4 (n=6) 9

of the current recorded before analyses. This was not unexpected, as Geobacter is 10

commonly exposed to metals in this potential range. Future work with stricter 11

anaerobes, or organisms adapted to higher-potential environments, should 12

independently verify the potential range which is not harmful to electron transfer. 13

Stable catalytic features with high signal-to-noise (S/N) ratio were observed 14

when CV was performed at slow scan rates (1 mV/s). Under these conditions, the rate 15

of electron transfer rose rapidly at a potential higher than –0.3 V, reaching a limiting 16

current above a potential of –0.1 V (Figure 4). This response, commonly called a 17

‘reversible catalytic wave’ (7), indicated continuous regeneration of the entire series of 18

proteins catalyzing electron transfer from the substrate to the electrode. In preliminary 19

investigations, each increase in scan rate significantly increased the ohmic current, and 20

shifted the location of peaks in voltammetric waves, as expected from kinetic effects 21

(Supplementary Figure S2). However, increasing scan rate five-fold did not significantly 22

alter the limiting current, demonstrating that at slow scan rates, conditions were within a 23

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timescale sufficient to establish sustained catalysis by cells at each applied potential. As 1

the most informative scan rate could vary for each experimental setup, scan rates used 2

in this report should be taken only as an aid in choosing conditions for new organism or 3

electrode surface studies. 4

The inflection point(s) in such catalytic waves were visualized as peaks using 5

first-derivative analysis of voltammetry data, (using software described in Methods) 6

(2,16). Such analysis of four independent biofilms is shown in Figure 4. One dominant 7

inflection point could be detected by this analysis, centered at about -0.15 V, which 8

increased in height with age of electrode-attached culture, while smaller intensity peaks 9

centered at -0.220 V and -0.020 V vs. SHE could also be detected. The dominant peak 10

revealed in derivative analyses was consistent with the presence of a single 11

oxidation/reduction event in electron transfer that was rate-limiting (1,9,15). As shown in 12

Figure 4, these values were highly repeatable between individual biological replicates. 13

Once a biocatalyst is adsorbed, it is also possible to determine its response to 14

increasing concentration of substrate (24). To test this approach, electron donor was 15

first removed from the medium, and electrodes were incubated at 0.24 V for 36 h, until 16

no residual anodic (oxidation) current was observed. At this point, the effect of lowering 17

substrate concentrations could be determined. Anodic limiting current at -0.1 V was 18

determined from CV, and was found to increase linearly with acetate concentration 19

when acetate was <200 µM. At acetate concentrations higher than 3 mM, anodic 20

limiting current did not change (Figure 5). The midpoint potential of catalytic waves 21

remained centered at the same position for all concentrations, suggesting a similar 22

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electron transfer mechanism and rate-controlling reaction, even at low current densities. 1

This dose-response test detected acetate at concentrations as low as 3.5 µM (S/N>3). 2

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Differential pulse voltammetry (DPV). Although CV is the most informative 4

method used to investigate catalytic substrate oxidation by adsorbed enzymes, it has a 5

low detection limit, and subtraction of ohmic (capacitive) current is necessary to reveal 6

small features (9). Furthermore, when electron transfer between adsorbed enzymes and 7

electrodes is slow, as is expected for complex electron transfer chains studied at 8

stationary electrodes, CV requires substantial time, and proper derivative analysis 9

requires post-processing of data (40). In comparison, pulse methods have the potential 10

to reveal characteristic peaks while cancelling out capacitive current, even at faster 11

scan rates, and are often used as complementary techniques to cyclic voltammetry 12

(20,44,65). 13

Preliminary experiments with G. sulfurreducens biofilms indicated that DPV could 14

also be used to monitor biofilms non-destructively, across a range of scan rates (up to 15

50 mV/s) and pulse heights (up to 100 mV). The parameters chosen (see Methods) 16

represent a compromise between analysis time and sensitivity. When performed on 17

mature biofilms, DPV always revealed a broad peak, which increased in height with the 18

age of the biofilm. These voltammograms were highly reproducible, with the peak 19

centered at -0.105±0.005 V vs. SHE (Figure 6). 20

Additional peaks at lower and higher potential with respect to the main peak were 21

also detected. The results were consistent with multiple redox centers exposed on the 22

surface of G. sulfurreducens cells that spanned a range of potentials (from 23

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approximately -0.2 to +0.1 V). In recent studies with Shewanella biofilms grown on 1

similar electrodes, we observed strikingly different behavior, as DPV revealed a large, 2

less convoluted peak at -0.21 V. These Shewanella cells were found to secrete redox-3

active flavins that mediated electron transfer (47), and changes in DPV peak height 4

could be used to monitor both the accumulation of this redox-active mediator at the 5

electrode and its loss when medium was changed to remove soluble compounds. As 6

DPV peak height was unaffected by medium changes in experiments with Geobacter 7

biofilms, this represented additional support for accumulation of an attached electron 8

transfer agent, and lack of soluble mediators within G. sulfurreducens biofilms. These 9

results demonstrate the utility of combining sweep and pulse methods in the study of 10

living systems. 11

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Electrochemical Impedance Spectroscopy (EIS). While originally applied to 13

investigation of solid-solid interfaces, EIS is today the most common alternating current 14

method applied to the characterization of aqueous biointerfaces (10,13). In most EIS 15

methods, a small sinusoidal potential perturbation is applied to the sample. The 16

frequency of this perturbation is changed in the range between few mHz and 105 Hz. 17

The resulting sinusoidal current is analyzed via Fast Fourier Transform techniques to 18

calculate the impedance (Z) of the interface in the frequency domain to estimate charge 19

transfer resistance (32), diffusion at surfaces covered by protein monolayers (20), 20

charge transfer time constants (59), and mechanisms of electron transfer (8). 21

Applications of EIS are numerous in microbially influenced corrosion, in which the 22

sample is studied at open circuit potential (42,51). However, we have shown that 23

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electron transfer processes by G. sulfurreducens are voltage-dependent (e.g. Figure 3). 1

Thus, while EIS is typically performed at open-circuit potentials, potentiostatic EIS 2

where the sinusoidal potential is applied while maintaining a given potential at the 3

working electrode was more likely to reveal electron transfer characteristics of active 4

cells attached to electrodes. 5

Figure 7 shows a comparison of a glassy carbon electrode before and after 6

colonization by G. sulfurreducens. When EIS was performed at V=-0.16 V vs. SHE, the 7

presence of G. sulfurreducens reduced charge transfer resistance (inferred from the Z 8

value at low frequencies), with a main time constant in the 1-10 Hz frequency range. At 9

high rates of electron transfer, a second, slower time constant was observed in the 10

0.01-0.1 Hz range. The addition of a second time constant to the model did not 11

significantly improve the dispersion of residuals, likely due to the low frequency of the 12

second time constant. 13

The importance of potentiostatic EIS was illustrated by the significant influence 14

that imposed potential had on charge transfer resistance and time constants for G. 15

sulfurreducens biofilms (Figure 8 and Table 1). When the imposed potential was similar 16

to the midpoint potential of the catalytic wave observed in CV, the lowest resistances 17

and time constants were measured. At potentials outside of this range, differences in 18

charge transfer were much smaller (in agreement with CV results), resulting in higher 19

apparent values for the charge transfer resistance and time constant. Across the 20

potential range explored, the solution resistance (R=17.2 ±0.4 Ohm), double-layer 21

capacitance (Cp=3.0±0.9 mF), and the nonideality coefficient (a=0.72±0.06) did not 22

change appreciably. These results again highlighted how data from one method (CV) 23

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could be used to select parameters in a complementary method (EIS). It also 1

demonstrated the importance of investigating the proper analysis parameters for each 2

bacterial system, as another strain with a different midpoint potential for its catalytic 3

process would need to be studied at a different imposed voltage to obtain the fastest 4

time constant. 5

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Implications. Many studies providing voltammetry data from bacteria-electrode 7

interactions have now been published. While reports differ in their pretreatment of 8

bacteria, biofilm age, growth medium, environmental conditions, electrode materials, 9

and electrochemical parameters, a general trend of catalytic wave-like behavior is clear 10

(14,31,53,68), showing that many bacteria and communities behave in a manner similar 11

to the early-stage Geobacter biofilms grown on geometrically simple electrodes shown 12

in this work. As a result, we can confirm what early observations suggested, that 13

Geobacter behave as attached biocatalysts, stimulated by changes across a narrow 14

potential range. For example, in a prior study, where we adsorbed washed Geobacter 15

cells on carbon paper electrodes (using pectin to retain free cells) (62), similar features 16

were reported even after a few hours of cells being exposed to electrodes. These 17

consistencies further suggest that these responses and potentials are not dramatically 18

influenced by such factors as stage of growth or electrode material. 19

In particular, this study shows how the response of an intact film to a range of 20

applied potentials can be measured systematically, and analyzed to produce data easily 21

compared between species, conditions, or electrode surfaces. As scan rates are 22

lowered, electron transfer kinetic effects (rates of electron transfer between cell surfaces 23

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and electrodes) are minimized, and enzymatic effects (related to microorganisms 1

activity) become primary factors. As the goal was to initially characterize the catalytic 2

behavior of the system, low scan rates (1 mV/s) were chosen, as they permitted 3

reactions with a time constant on the order of ~1 s to be active as turnover processes at 4

each imposed potential step. At high levels of electron donor, and low scan rates, 5

catalytic voltammograms should therefore be representative of steady-state conditions. 6

In addition, minimization of ohmic current aided in identification of inflection points in 7

derivative analysis. The choice of a relatively low scan rate was also supported by data 8

from EIS measurements, in which we found time constants on the order of 0.1 and 1-10 9

s. For a system such as this, with what appears to be sluggish interfacial electron 10

transfer kinetics, the potential difference between anodic and cathodic peaks for a given 11

redox couple could change significantly with even modest changes in scan rate. Future 12

work will focus on the use of electrochemical techniques under non-turnover conditions, 13

to better elucidate these electron transfer kinetics, for a more complete understanding of 14

the interplay between microbial catalytic abilities and interfacial electron transfer. 15

The high reproducibility between biological replicates and ability to perform 16

experiments with a range of electrodes provided a robust measurement of G. 17

sulfurreducens current densities, which approached 2 A/m2 for electrodes polished with 18

35 µm grit size particles. The catalytic wave consistently observed for G. sulfurreducens 19

is an independent demonstration that interior oxidative processes of this organism are 20

linked via a continuous pathway to surfaces, and that the entire collection of attached 21

organisms (i.e., the biofilm) behaves as an adsorbed catalyst. The midpoint potential of 22

the catalytic wave at -0.15 V supports a model with a dominant rate-limiting electron 23

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transfer reaction, and shows that G. sulfurreducens respiration rate does not increase 1

when cells are provided with an electron acceptor with a potential greater than 0 V. The 2

latter implies that the final step of electron transfer (e.g. between a terminal external 3

protein and the electrode) is not rate-limiting, as this process can always be accelerated 4

by additional applied potential (as described by the standard rate law kf=k0 eß∆E (9)). 5

The midpoint and the limiting current potential found in this study are consistent with G. 6

sulfurreducens being adapted for the reduction of iron oxides with potential between 7

-0.2 and 0 V vs. SHE in the environment, and suggest that cells do not derive any 8

additional energetic benefit from higher-potential electron acceptors. 9

While the study of an intact electron transport chain in G. sulfurreducens 10

represents perhaps a complex example of this electrochemical approach, similar 11

methods have been used to characterize extracts from bacterial cells adsorbed onto, or 12

incorporated into, electrode materials (26,45,46,61). In addition, researchers have 13

washed and concentrated whole cells for brief voltammetry analyses and detected 14

possible redox-active agents on the external surface of cells (39). The benefits of using 15

living cells in the work reported here include a lack of a protein purification step, and the 16

fact that cells are allowed to assemble the actual biofilm structure responsible for this 17

unique extracellular electron transfer process. 18

The multiple peaks visible in both CV and DPV analyses also confirm the 19

complexity of the G. sulfurreducens surface. Multiple cytochromes and redox proteins 20

have been previously implicated in outer membrane-based electron transfer in 21

proteomic and labeling studies (23,36,41). As most proteins implicated in electron 22

transport by G. sulfurreducens contain multiple hemes or redox centers, the detected 23

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redox centers could reflect individual hemes, domains that act as a single center, or 1

individual proteins. Recent work with the multiheme cytochrome MtrC (30) showed that 2

multiheme proteins do not demonstrate classic, individual redox behavior for each 3

heme, but rather act as a cluster with a broader midpoint. In another study (67), the 4

same protein was observed to behave as two pentaheme domains with broad midpoint 5

potentials. Future work with specific mutants lacking key redox proteins in G. 6

sulfurreducens will aid in identifying the origin of these peaks. 7

Based on the results reported here, voltammetric methods previously developed 8

to characterize electron transfer phenomena by enzymes adsorbed at carbon 9

electrodes can be extended to the characterization of viable biofilms. By appropriate 10

choice of the conditions, these methods are not destructive and allow in vivo 11

determination of electron transfer from whole cells to electrodes under conditions that 12

are comparable to those encountered in natural environments. Both thermodynamic and 13

kinetic parameters can be determined and used to define the phenotype of an organism 14

for comparison with other strains or mutants. The proposed methods can be applied to 15

well-defined pure cultures, as well as to complex microbial communities, and could 16

allow for quantitative comparisons in the development of better microbial catalysts 17

based on direct electron transfer between bacteria and electrodes. 18

19

ACKNOWLEDGEMENTS 20

We thank IREE and NSF (Grant DBI-0454861) for their support, as well as the 21

BioTechnology Institute at University of Minnesota. We also thank Jeremy Alley for 22

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technical help with the experimental setup, Dr. Dirk Heering for suggestions in the 1

interpretation of data, and Lewis Werner for the SEM images of bare electrodes. 2

3

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Figure legend 1

Figure 1 : (A) Cartoon of 3-electrode bioelectroreactor and (B) connection scheme to 2

multichannel potentiostat. 3

Figure 2: Characteristic growth curve for G. sulfurreducens inoculated into a bioreactor 4

containing a 2.5 cm2 glassy carbon electrode (P400 grit polishing). ‘Analysis’ breaks 5

indicate when cyclic voltammetry and differential pulse voltammetry was performed. 6

‘Medium change’ breaks indicate where planktonic cells and culture medium was 7

removed and replaced with fresh medium. Raw data is shown with no smoothing or 8

noise-reduction transformations. 9

Figure 3: SEMs of glassy carbon polished with P400 vs. P4000 grit treatment (top). 10

Bar graph (below) shows maximum current measured (n=2 for each roughness) during 11

chronoamperometry, after 72 hours of growth. Also indicated ((), right Y axis) is the 12

average grit size of each polishing treatment. 13

Figure 4: (A) Low scan rate cyclic voltammetry of G. sulfurreducens biofilms after 14

inoculation (0 h) and at maximum current production (72h). (B) First derivatives of cyclic 15

voltammograms of biofilms (P400 grit polishing) (n=4) showing the midpoint potential 16

detectable in catalytic waves of mature biofilms. 17

Figure 5: Low scan rate cyclic voltammetry (1 mV/s) of G. sulfurreducens at different 18

concentrations of electron donor. The biofilm was poised at oxidizing potential without 19

electron donor until no anodic current was observed. Then electron donor was added 20

and the limiting current was recorded (inset). The midpoint potential of catalytic curves 21

was constant with increasing concentration of electron donor. 22

23

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Figure 6: (A) Differential pulse voltammetry (SR 4 mV/s, PH 50 mV) under catalytic 1

(presence of acetate) conditions at inoculation and after 72 h of growth. (B) The shape 2

of peaks and average potential for DPV of mature biofilms (72 h) between 4 3

independent biological replicates. 4

Figure 7: EIS of G. sulfurreducens biofilms at V=-0.16 V vs. SHE: modulus of 5

impedance Z after inoculation () and at 72 h (); phase angle V-I after inoculation () 6

and at 72 h (). Biofilm development results in lower charge transfer resistance at the 7

electrode-biofilm interface, with two distinct charge transfer processes around 1 and 8

0.03 Hz. 9

Figure 8: EIS of mature G. sulfurreducens biofilms (72 h) performed at different 10

potentials: (A) Bode plot. Modulus of impedance Z at 0.04 (black full circles), -0.06 (red 11

full circles), -0.16 (blue full circles), and -0.26 V (green full circles) vs. SHE; phase 12

angle V-I at 0.04 (black empty circles), -0.06 (red empty circles), -0.16 (blue empty 13

circles), and -0.26 V (green empty circles) vs. SHE. Note that frequency of the major 14

charge transfer process changes with applied potential. The second charge transfer 15

process can be observed only at -0.16 V. (B) Nyquist plot. The relaxation times for the 16

electron transfer process at the interface are summarized in Table 1. 17

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Table 1: EIS parameters for biofilms at 72h. Data were fitted with the single time 1

constant model described in the methods. 2

Potential vs. SHE

(Volts)

Charge transfer

resistance (Ohm)

Characteristic

frequency (Hertz)

Time constant (sec)

+0.042 23500 <0.01 >100

-0.058 1050 0.032 32

-0.158 204 0.5 2

-0.258 2440 0.04 25

3

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