Electron transfer and biofilm formation of Shewanella putrefaciens as function of anode potential

Bioelectrochemistry xxx (2012) xxx–xxx

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Bioelectrochemistry

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Electron transfer and biofilm formation of Shewanella putrefaciens as function ofanode potential

Alessandro A. Carmona-Martínez a, Falk Harnisch a,⁎, Ute Kuhlicke b, Thomas R. Neu b, Uwe Schröder a

a Institute of Environmental and Sustainable Chemistry, Technische Universität Braunschweig, Hagenring 30, 38106 Braunschweig, Germanyb Helmholtz Centre for Environmental Research UFZ, Department of River Ecology, Brückstraße 3a, D-39114 Magdeburg, Germany

⁎ Corresponding author. Tel.: +49 531918428; fax: +E-mail address: [email protected] (F. H

1567-5394/$ – see front matter © 2012 Elsevier B.V. Alldoi:10.1016/j.bioelechem.2012.05.002

Please cite this article as: A.A. Carmona-Mapotential, Bioelectrochemistry (2012), doi:1

a b s t r a c t
a r t i c l e i n f o
Article history:Received 27 January 2012Received in revised form 23 April 2012Accepted 3 May 2012Available online xxxx

Keywords:Microbial bioelectrochemical systemsMicrobial fuel cellsBacterial electron transferShewanella putrefaciensCyclic voltammetry

Shewanellaceae are among the most widely studied electroactive microorganisms. In this report, we studied theinfluence of the applied electrode potential on the anodic current production of Shewanella putrefaciens NCTC10695 under anoxic conditions. Furthermore, we used cyclic voltammetry (CV) and confocal laser scanning mi-croscopy (CLSM) to investigate the microbial electron transfer and biofilm formation. It is shown that the chro-noamperometric current density is increasing with increasing electrode potential from 3 μA cm−2 at−0.1 V upto ~12 μA cm−2 at +0.4 V (vs. Ag/AgCl), which is accompanied by an increasing amount of biomass depositedon the electrode. Bymeans of cyclic voltammetrywe demonstrate that direct electron transfer (DET) is dominat-ing and the planktonic cells play only a minor role.

© 2012 Elsevier B.V. All rights reserved.

1. Introduction

Although, the ability of microorganisms to create an electrochemicalpotential was discovered more than 100 years ago [1], it was mainlyduring the last decade that this field of research was developing mostdramatically [2], as itwas found that the ability to transfer electrons to ex-tracellular electron acceptors is naturally occurring in several microbialspecies. This interest thereby is triggered by fundamental inquisitiveness[3–5] as well as by the development of seminal sustainable technologiesbased on microbial extracellular electron transfer—also known as micro-bial bioelectrochemical systems (BES) [6,7].

Up to now, severalmicroorganisms have been studied in order to elu-cidate their specific extracellular electron transfermechanisms, includingLactococcus lactis [8], Saccharomyces cerevisiae [9], Pseudomonas sp.CMR12a [10], Hansenula anomala [11], Proteus vulgaris [12] and Lyngbyasp. and Nostoc sp. [13]. Here, the two families of Gram-negative bacteriaShewanellaceae [14,15] and Geobacteraceae [16–19] are the most wide-spread model organisms. Thereby, different electron transfer strategieshave been commonly described among the various species, which aresummarized as follows: i) direct electron transfer (DET) via membrane-bound redox-enzymes [20–22] or via bacterial nanowires [17,23–25]and ii) mediated electron transfer (MET) via redox shuttle substances,e.g. [26,27], usually secondary microbial metabolites.

49 531918424.arnisch).

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rtínez, et al., Electron transfer0.1016/j.bioelechem.2012.05.

For the family of Shewanellaceae – all being considered to be faculta-tive anaerobes – several members have been studied on their extra-cellu-lar electron transfer behavior, most prominently Shewanella oneidensisMR-1 [28–31], but also S. oneidensis MR-4 [27], Shewanella putrefaciensW3-18-1 [32], S. putrefaciens IR-1 [33–35], S. putrefaciens SR-21 [33],Shewanella loihica PV-4 [32,36–39], Shewanella decolorationis NTOU1[40,41], Shewanella japonica KMM 3299 [42], Shewanella frigidimarinaNCIMB400 [43,44] and Shewanella marisflavi EP1 [45]. Commonly, it is as-sumed that the two decaheme c-type cytochromesMtrC and OmcA, bothfacing the extracellular environment, play a key role in the direct electrontransfer mechanism (DET) [28,46–50]. Thereby, MtrC and OmcA, are partof a complex transmembrane cytochrome pool involving more than 40proteins [51].

The MET of Shewanellaceae exploits redox-shuttles including humicsubstances [52], melanin [53,54], menaquinone [55] as well as riboflavin[56],flavinmononucleotide [42,57] and their derivatives. Further it is con-sidered to depend on intracellular electron transfer to the redox-shuttleas well as extracellular electron transfer by MtrC or OmcA, e.g. Refs.[27,42,58]. Thereby it has been shown that some strains of the Shewanellafamily (e.g. S. oneidensis MR-1 [27,59], S. loihica PV-4 [60], Shewanellabaltica Os155 and Os195 [56], S. frigidimarina NCIMB400 [56] andS. decolorationis NTOU1 [40]) can biosynthesize these mediators underaerobic as well as anaerobic cultivation. However, the ability to synthe-size suitable amounts of these electron shuttles (especially for anaerobicconditions, where energy for biosynthesis is limited) is not unequivocallyestablished for all Shewanella species, e.g. Ref. [56]. In this course it is ofcrucial importance to consider the exact conditions during biofilmgrowth and development [61].

and biofilm formation of Shewanella putrefaciens as function of anode002

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2 A.A. Carmona-Martínez et al. / Bioelectrochemistry xxx (2012) xxx–xxx

1.1. Influence of the electrode potential on electroactive microbial biofilms

It has been shown for anodic mixed culture derived biofilms thatthe applied electrode potential (denominated as Eappl within this arti-cle) determines decisively the bacterial composition, e.g. Ref. [62], ofthe gained biofilm. However, also pure culture anodic biofilms andthe microorganisms therein can be influenced by the applied elec-trode potential. Cho and Ellington [63] demonstrated for S. oneidensisMR-1 that during chronoamperometric biofilm growth the lag-perioddecreased from 90 days at 0 mV to 5 days at 500 mV (vs. Ag/AgCl), re-spectively with increasing electrode potential, whereas the currentdensity was almost constant. Furthermore, Shewanella oneidensisMR-1 is believed to show some motility towards electrodes andthus the availability of its cellular appendices (wires, pili and flagella)clearly determines its electron transfer performance [64]. RecentlyHarris and colleagues [65] observed an increase in cell swimmingspeed when whole cells of Shewanella species (S. oneidensis MR-1,Shewanella amazonensis SB2B and S. putrefaciens CN32) were exposedto varying electrode potentials. In addition, Liu and colleagues [66]investigated microbial extracellular electron transfer activity ofS. loihica PV-4 by applying different electrode potentials (from −0.4 Vto 0.2 V vs. Ag/AgCl). Thereby, they demonstrated a clear dependenceof the activity on the applied electrode potential, with an activity in-crease for potentials more positive than −220 mV. Furthermore, Liuet al. [67] have recently shown on the example of S. oneidenis MR-1and S. loihica PV-4 that, depending on the potential of bacteria cultiva-tion, the electron transfer pathways can be switched from DET to METand the formal potential of the DET [66] is also triggered by theredox-conditions during cultivation.

Within the context of correlating the applied electrode potential (i.e.potential of the microbial terminal electron acceptor) and the bacterialactivity, it was the aim of this study to elucidate the electrochemical re-sponse of S. putrefaciens NCTC 10695, a strain that is studied in BES forthe first time. (For a comparison with other S. putrefaciens see TableS2.) Therefore, the influence of the applied electrode potential on theelectron transfermechanismswas investigated using cyclic voltammetry(CV) and biofilm morphology by means of confocal laser scanning mi-croscopy (CLSM). CLSM has recently been applied for the study of elec-troactive biofilms, e.g. to monitor their pH-gradients [68], but has notyet been exploited for the quantification of biofilm biomass for differentelectrode potentials.

2. Materials and methods

2.1. General conditions

All chemicals were of analytical or biochemical grade and werepurchased from Sigma-Aldrich and Merck. If not stated otherwise,all potentials provided in this article refer to the Ag/AgCl referenceelectrode (sat. KCl, 0.195 V vs. SHE). All microbial experiments wereperformed under strictly sterile conditions.

2.2. Cell cultures and media

Sterilized LB broth [69], and minimal media [70] (18 mmol/L sodiumlactate, PIPES buffer 15.1 g×L−1; NaOH 3.0 g×L−1; NH4Cl 1.5 g×L−1;KCl 0.1 g×L−1; NaH2PO4∙H2O 0.6 g×L−1; NaCl 5.8 g×L−1; Wolfe's min-eral solution 10 mL×L−1 [71];Wolfe's vitamin solution 10 mL×L−1 [71];amino acid solution 10 mL×L−1 [70]) for liquid cultures and LB/agar(Merck KGaA, Germany) plates were used for culture maintenance.

Single colonies on LB/agar plates freshly streaked from a frozenglycerol stock culture [72] of S. putrefaciens wild-type NCTC 10695[73–76] (DSM No.: 1818, DSMZ—German Collection of Microorgan-isms and Cell Cultures GmbH, Braunschweig, Germany) were trans-ferred to 15 mL of LB broth and incubated aerobically at roomtemperature while shaking at 100 rpm (Universal shaker SM 30 A,

Please cite this article as: A.A. Carmona-Martínez, et al., Electron transferpotential, Bioelectrochemistry (2012), doi:10.1016/j.bioelechem.2012.05.

Edmund Bühler GmbH, Germany) for 48 h [64]. Afterwards 15 mL ofculture were spun down at 3000 rpm during 10 min (HeraeusMegafuge 1.0, Germany). The pellet was resuspended in 15 mL ofminimal media and transferred to a sterile Erlenmeyer flask with185 mL of minimal media for 72 h of cultivation using the same con-ditions. Finally 200 mL of media were centrifugated at 3000 rpm dur-ing 10 min; the pellet was resuspended in 15 mL of minimal mediaand injected in the electrochemical cell.

2.3. Bioelectrochemical set-up and experiments

Polycrystalline carbon rod (CPGraphite GmbH, Germany)was used asworking (2.0 cm height×1.0 cm diameter) and as counter (7.0 cmheight×1.0 cm diameter) electrodes for the growth and investigation ofthe (anodic) electrocatalytic biofilms. Carbon electrodes were gluedwith stainless steel wire (AISI 304, Fe/Cr18/Ni10, Goodfellow GmbH,Nauheim, Germany) using a two-component resin (Epoxy resin HT 2+Hardener HT 2, HY-POXY® Systems, SC, USA) mixed with carbon blackparticles (Vulcan XC-72, Cabot Corporation GMBH, Frankfurt am Main,Germany).

All bioelectrochemical experiments were conducted under strictlysterile conditions and under potentiostatic control, using Ag/AgCl refer-ence electrode (sat. KCl, 0.195 V vs. SHE, Sensortechnik Meinsberg,Germany). S. putrefaciens biofilms were grown in potentiostatic half-cellexperiments at six different anode potentials (−0.1, 0, +0.1, +0.2,+0.3 and +0.4 V) at 30 °C [77]—see also Figure S1. To assure compara-bility and reproducibility up to six electrodes were measured simulta-neously in one electrochemical cell using an Autolab 30 potentiostat(Ecochemie, TheNetherlands) equippedwith six array channels. The bio-film growth was performed in semi-batch chronoamperametric experi-ments at different applied potentials with a regular Shewanella cellsaddition procedure [29] and media replacement as described below. Forthese potentiostatic biofilm growth experiments, initial aerobic condi-tions were assured according to previous reports [78]. Therefore, beforeadding the growth media into the electrochemical cell [56,79] filteredair was pumped into the fresh minimal media using a fish pump (Eliteair pump 799, Rolf C. Hagen Corp., MA, USA) per 400 mL for more than1 h. Fresh electron donor and nutrients were supplied about every 72 hby removing 320 mL of culture (representing 80% of the cell volume)and replacing with fresh minimal media and fresh Shewanella cellsaccording to Section 2.2. Cyclic voltammograms were recorded for turn-over and non-turnover conditions according to Ref. [64].

2.4. Electrochemical data processing

All data are based on experiments of 3 independent sets of biofilmreplicates and the standard deviations are presented. The maximumcurrent density, jmax, (calculated per projected electrode surfacearea) of a batch-cycle was taken as a measure of activity and was an-alyzed during at least 6 semi-batch cycles for established microbialbiofilms. For in-depth data analyses of the cyclic voltammogramsthe open‐source software SOAS [80] was used for baseline (capacitivecurrent) correction for non-turnover conditions.

2.5. Confocal laser scanning microscopy

Shewanella biofilms on polycrystalline graphite electrodes wereexamined by CLSM after staining with nucleic acid-specific fluoro-chromes. For this purpose, whole cylindrical electrodes (~2.0 cm inexposed length) were mounted on a plastic Petri dish with siliconglue, subsequently stained with Syto9 (Molecular Probes, OR, USA)and incubated at room temperature for 5 min [81]. The laser micro-scope (SP5X, Leica Germany) was equipped with a super continuumlight source and an upright microscope. The system was controlledby the LEICA CONFOCAL software version 2.4.1 Build 1537 (Leica,Germany). For imaging, the stained biofilms were covered with tap



3A.A. Carmona-Martínez et al. / Bioelectrochemistry xxx (2012) xxx–xxx

water and examined with a 63×0.9 NA objective lens. EachShewanella biofilm attached to the electrode was scanned at ten dif-ferent locations from top to bottom. The settings for excitation andemission/detection were as follows: excitation with the white laserat 483 nm, detection of reflection from 475 to 495 nm and of Syto9from 500 to 570 nm. For recording the bacterial emission signal, thelookup table “glow-over-under”was used in order to optimally adjustsignal to noise ratio and taking advantage of the full dynamic range ofthe photomultiplier. Quantification was done with an extended ver-sion of ImageJ (rsbweb.nih.gov.ij) as described elsewhere [82]. The8 bit data sets were thresholded at 60. Image data sets were printedfrom Photoshop (Adobe).

Fig. 2. Chronoamperometric current density of Shewanella putrefaciens as function ofthe applied electrode potential during chronoamperometric growth (Eappl).

3. Results and discussion

3.1. Bioelectrochemical current production

Fig. 1 shows for exemplary fed-batch cycles the chronoamperometric(CA) current production of S. putrefaciens NCTC 10695 for different ap-plied electrode potentials. As typical for fed-batch operation the currentincreases after media replenishment to a maximum, jmax, and decreasesthereafter due to substrate exhaustion. As can be seen themaximumcur-rent density, jmax, as well as the transferred charge of each electrode dif-fered significantly. Since all electrodes were hosted within one vessel inorder to assure similar biological conditions for all electrodes in each ex-periment (see Material and methods), no coulombic efficiencies can becalculated for the individual electrode potentials.

However, when taking jmax at a given electrode potential a cleartrend can be observed. As Fig. 2 shows the current density increasesfrom less than 3 μA cm−2 at −0.1 V up to ~12 μA cm−2 at +0.4 V.

Thus, the measured current densities are well in line with previ-ous studies on Shewanellaceae using plain carbon electrodes (seeTable 1 in SI for a comparative data compilation). No biofilms couldbe seen by the naked eye on all electrodes, but the solution showedsignificant turbidity and a reddish pellet of bacteria was formedwhen spinning it down (see Fig. S1—inset). This finding is well in ac-cordance with previous reports on Shewanaellaceae [31,68,72,83], yetin contrast to pure culture biofilms of Geobacter species or mixed cul-ture derived biofilms dominated by Geobacteraceae. In order to eluci-date reasons for the varying electrochemical performance at thedifferent electrode potentials, cyclic voltammetry (CV) and confocallaser scanning microscopy (CLSM) were employed.

Fig. 1. Representative chronoamperometric fed-batch cycles of Shewanella putrefaciensat graphite electrodes; applied potentials: a) −0.1; b) 0; c) +0.1; d) +0.2; e) +0.3and f) +0.4 V vs. Ag/AgCl; CV measurements during turn-over (A) and non turn-over (B) conditions respectively.


3.2. Cyclic voltammetric analysis

Fig. 3 shows the CVs of the electrodes under turn-over conditions,i.e. in the presence of the electron donor lactate. All CVs show an in-creasing bioelectrocatalytic activity for all potentials more positivethan 0.0 V and possess a half wave potential of about ~10 mV (ascan be derived from the maxima of the respective first derivatives[84,85]). This potential can commonly be ascribed to DET-proteins(see e.g. Refs. [49,86]). In this course it has to be noticed that thisis the first voltammetric study of S. putrefaciens NCTC 10695 (forS. putrefaciens from other sources see Table S2).

Thus, the results for turn-over conditions point out that the MET viasolublemediators plays only aminor role, as the formal potentials, Ef, forthese electron shuttling compounds (and thus the corresponding halfwave potential of the turn-over CV) are more negative. Noteworthy,the riboflavin concentration of our minimal media nutrient broth wasonly about 0.1 μM (see Materials and methods). When this concentra-tionwas increased to 1 μM, the turn-over CV-shape changed significant-ly and clearly illustrated that the riboflavin based MET was dominant inthese conditions (see SI Figures S2 and S3). Thus, as no significant METassociated bioelectrocatalysis was found after more than 2 weeks of

Fig. 3. Representative cyclic voltammograms of S. putrefaciens for turn-over conditionsand B) respective first derivatives of the voltammetric curves; scan rate of 1 mV s−1.




continuous cultivation, our results suggest that S. putrefaciens NCTC10695 did not synthesize relevant amounts of electron shuttle com-pounds. This finding is in contrast to other species of the Shewanellafamily like S. oneidensisMR-1 [27,56,59].We suggest to ascribe this find-ing to the particular growth conditions especially i) the almost anoxicbehavior and ii) low mediator concentration in the medium and thusfast establishment of a biofilm at the electrode.

In order to study the biofilm associated direct electron transferprocesses in more detail, CVs for non turn-over (lactate depleted)conditions were recorded (see Fig. 4 A for the raw data and Fig. 4 Bfor the baseline corrected traces). It can be clearly seen that the nonturn-over CVs for the selected electrode potentials differed signifi-cantly. As all electrodes were hosted within one vessel, these differ-ences cannot be attributed to the suspended microbial cells, but tothe individual biofilms. Noteworthy, no CV signals were detected forelectrodes poised at −0.1 V and 0.0 V during biofilm growth, whichcan clearly be attributed to the absence of the interaction with bacte-ria on the electrode surface (vide infra). As Fig. 4 shows, the CV anal-ysis of all further electrodes revealed that the formal potential of theredox-active species is similar for all growth potentials (about −60 mV, see also Table S3). This value is close to the maximum in thefirst derivatives of the turn-over CVs (Fig. 3) showing that thedetected redox couple is responsible for the bioelectrocatalytic elec-tron transfer.

As can be seen from Fig. 4 the peak heights and areas of the oxida-tion and reduction peaks are increasing with more positive electrodepotential during biofilm growth.

Deriving from the data exemplary shown in Fig. 4, Fig. 5 depicts thepeak areas and peak heights of the respective oxidation and reductionpeak as function of the applied potential, Eappl, during CA. As the peakarea can be considered to be ameasure of the amount of a redox specieson the electrode surface [87,88], this analysis shows that with increas-ingly positive potential during bacterial growthmoreDET-protein is de-posited on the electrode surface. As the increasing amount of DETprotein can be either attributed to an increasing cell number or to an in-crease in the protein amount per individual cell confocal laser scanningmicroscopy (CLSM) was exploited to study these biofilms at the mor-phological level.

Fig. 4. A) Cyclic voltammograms for non turn-over conditions for Shewanella putrefaciensusing a scan rate of 1 mV s−1; B provides the respective baseline corrected curves. For A)and B); applied potentials during chronoamperometric growth: a) +0.1; b) +0.2; c)+0.3 and d) +0.4 V vs. Ag/AgCl.


As already discussed above, the detected formal potential is close tobut not identical to those of MtrC and OmcA commonly reported in lit-erature for different experimental conditions, e.g. on isolated proteins[49]. Here it should be noticed that additional cyclic voltammetry ex-periments conducted on S. putrefaciens cells suspensions (data notshown) did not provide redox peaks in any condition tested, i.e. 1) bac-teria growing microaerophilically in LB broth, 2) cells growing aerobi-cally in minimal media [70] and 3) cells growing anaerobically in [89]minimal media [27,28,70] using sodium fumarate as electron acceptor.

3.3. Biofilm imaging using confocal laser scanning microscopy (CLSM)

In order to study the biofilm formation at the electrode surface ex-emplary CLSM measurements were performed. As Fig. 6 shows thenumber of cells [90] seems to increase with more positive electrodepotential. This is confirmed by a biomass quantification based onthe CLSM imaging and subsequent digital image analysis (Fig. 7).Please note that the high numeric value in Fig. 7 for −0.1 V canonly be attributed to a heterogenous biofilm growth and biofilmsloughing. This effect needs further consideration in future studieswith focus on biofilm development over time. The data are showinga linear increase of biomass with more positive electrode potential.Nevertheless the relatively high standard deviation is obvious andwell known from other studies, e.g. Ref. [91], indicating a non-uniform (rough) biofilm coverage. Interestingly, as Fig. 6 shows, nocomplete coverage of the electrode surface and decent biofilm forma-tion was detected for all applied electrode potentials. This finding iswell in line with previous studies on Shewanella [31,68,72,83], butin clear contrast to Geobacter species [90,92]. Therefore, and in combi-nation with the dominance of DET and planktonic cell growth, onemay conclude that attachment of S. putrefaciens cells was not perma-nent, indicating an intermittent cell-electrode contact for electron re-lease [65]. Furthermore, this shows clearly the attraction of the morepositive potential for biofilm formation, which is in line with a reporton the bacterial movement to electrodes, termed “electrokinesis”[65].

4. Conclusion and outlook

This study on the electroactive microorganism S. putrefaciensNCTC 10695 shows the following:

• The chronoamperometric current generation, jmax, is a function of theapplied electrode potential; it increases from 3 μA cm−2 at−0.1 V upto ~12 μA cm−2 at +0.4 V.

• The amount of biomass deposited on the electrode is a function of theelectrode potential and steadily increase by factor 5 from0 V to+0.4 V.

• For the range of applied electrode potentials studied the direct elec-tron transfer (DET) is dominating.

Fig. 5. Plot of the base line corrected height (jp) and area (Qp) of the oxidation and re-duction peaks of redox-system shown in Fig. 4 as function of the applied potential dur-ing chronoamperometric growth. For visual convenience, reduction peak areas areshown as negative values. Symbols for peak height (○) and peak area (□).



Fig. 6. Maximum intensity projection of confocal laser scanning microscopy data sets showing Shewanella putrefaciens biofilms grown on electrode surfaces at different applied po-tentials. A) −0.1 V, B) 0 V, C) +0.1 V, D) +0.2 V, E) +0.3 V and F) +0.4 V; (all vs. Ag/AgCl). Color allocation: reflection of electrode—gray, nucleic acid stained bacteria—green.

Fig. 7. Volumetric biofilm quantification of Shewanella putrefaciens biofilms grown onelectrode surfaces at different applied potentials during chronoamperometric growth;ρbiomass provided in µm3

biomass per µm2electrode. Bacterial biomass was determined by

laser microscopy imaging of electrode surfaces with subsequent digital image analysisof 3d data sets (see materials and methods for details).



• The bioelectrocatalytic current generation, the CV signal of the DETprotein and the biomass coverage are clearly linked by the appliedelectrode potential during biofilm growth.

Interestingly, for electrode potentials more positive than the for-mal potential of the active site (−60 mV) the current density andbiofilm formation increase. The same phenomena are found for theturn-over CVs not showing a plateau of current density, which is incontrast to species like Geobacter [84].

Most of the results found for S. putrefaciens NCTC 10695 are well inline with literature; however some vary. These variations might be at-tributed to i) the specific nature of the strain S. putrefaciens NCTC10695 or ii) the specific growth conditions and thus may change inother environments, e.g. presence vs. absence of O2 in the mediumor constant vs. varying electrode potentials. Therefore, they are in arow of studies on Shewanella species, providing snapshots but notallow drawing conclusive pictures. Thereby, this study highlights ex-emplary the need for a unified framework of biofilm growth and




operation of electroactive biofilms, like Shewanellaceae [61] in order to ex-tract more (universal) information out of the gained data. This will cer-tainly be needed to shed light on the fundamentals of bacterial electrontransfer and to pave the way to application of microbial bioelectro-chemical systems.

Acknowledgments

The authors thank LucianoD. Dantas for helpwith the Linux and SOASsoftware handling. A.A.C.-M. thanks theDAAD for a Ph.D. scholarship. F.H.acknowledges the support by the Fonds der Chemischen Industrie (FCI).U.S. acknowledges the foundation of theprofessorship Sustainable Chem-istry and Energy Research by the Volkswagen AG and the Verband derDeutschen Biokraftstoffindustrie e.V.

Appendix A. Supplementary data

Supplementary data to this article can be found online at http://dx.doi.org/10.1016/j.bioelechem.2012.05.002.

References

[1] M.C. Potter, Electrical effects accompanying the decomposition of organic com-pounds, Proc. R. Soc. A 84 (1911) 260–276.

[2] U. Schröder, Discover the possibilities: microbial bioelectrochemical systems and therevival of a 100-year–old discovery, J. Solid State Electrochem. 15 (2011) 1481–1486.

[3] M.E. Hernandez, D.K. Newman, Extracellular electron transfer, Cell. Mol. Life Sci.58 (2001) 1562–1571.

[4] U. Schröder, Anodic electron transfer mechanisms in microbial fuel cells and theirenergy efficiency, Phys. Chem. Chem. Phys. 9 (2007) 2619–2629.

[5] K. Watanabe, M. Manefield, M. Lee, A. Kouzuma, Electron shuttles in biotechnol-ogy, Curr. Opin. Biotechnol. 20 (2009) 633–641.

[6] K. Rabaey, L. Angenent, U. Schröder, J. Keller (Eds.), Bioelectrochemical Systems: fromExtracellular Electron Transfer to Biotechnological Application, IWA Publishing,London, 2010.

[7] K. Rabaey, R.A. Rozendal, Microbial electrosynthesis—revisiting the electricalroute for microbial production, Nat. Rev. Microbiol. 8 (2010) 706–716.

[8] M. Masuda, S. Freguia, Y.-F. Wang, S. Tsujimura, K. Kano, Flavins contained inyeast extract are exploited for anodic electron transfer by Lactococcus lactis, Bio-electrochemistry 78 (2010) 173–175.

[9] R. Ducommun, M.-F. Favre, D. Carrard, F. Fischer, Outward electron transfer bySaccharomyces cerevisiae monitored with a bi-cathodic microbial fuel cell-typeactivity sensor, Yeast 27 (2010) 139–148.

[10] T. Pham, N. Boon, P. Aelterman, P. Clauwaert, L. De Schamphelaire, L. Vanhaecke,K. De Maeyer, M. Höfte, W. Verstraete, K. Rabaey, Metabolites produced by Pseu-domonas sp. enable a Gram-positive bacterium to achieve extracellular electrontransfer, Appl. Microbiol. Biotechnol. 77 (2008) 1119–1129.

[11] D. Prasad, S. Arun, M. Murugesan, S. Padmanaban, R.S. Satyanarayanan, S. Berchmans,V. Yegnaraman, Direct electron transfer with yeast cells and construction of amediatorless microbial fuel cell, Biosens. Bioelectron. 22 (2007) 2604–2610.

[12] F.J. Rawson, D.J. Garrett, D. Leech, A.J. Downard, K.H.R. Baronian, Electron transferfrom Proteus vulgaris to a covalently assembled, single walled carbon nanotube elec-trode functionalised with osmium bipyridine complex: application to a whole cellbiosensor, Biosens. Bioelectron. 26 (2011) 2383–2389.

[13] J. Pisciotta, Y. Zou, I. Baskakov, Role of the photosynthetic electron transfer chain inelectrogenic activity of cyanobacteria, Appl. Microbiol. Biotechnol. (2011) 377–385.

[14] C.R. Myers, K.H. Nealson, Bacterial manganese reduction and growth with manga-nese oxide as the sole electron acceptor, Science 240 (1988) 1319–1321.

[15] K.H. Nealson, J. Scott, Ecophysiology of the genus Shewanella, in: M. Dworkin,S. Falkow, E. Rosenberg, K.H. Schleifer, E. Stackebrandt (Eds.), The Prokaryotes,Springer Science, Singapore, 2006, pp. 1133–1151.

[16] F. Caccavo Jr., D.J. Lonergan, D.R. Lovley, M. Davis, J.F. Stolz, M.J. McInerney,Geobacter sulfurreducens sp. nov., a hydrogen- and acetate-oxidizing dissimilatorymetal-reducing microorganism, Appl. Environ. Microbiol. 60 (1994) 3752–3759.

[17] N.S.Malvankar,M.Vargas, K.P. Nevin, A.E. Franks, C. Leang, B.-C. Kim,K. Inoue, T.Mester,S.F. Covalla, J.P. Johnson, V.M. Rotello, M.T. Tuominen, D.R. Lovley, Tunable metallic-likeconductivity in microbial nanowire networks, Nat. Nanotechnol. 6 (2011) 573–579.

[18] H. Richter, K.P. Nevin, H. Jia, D.A. Lowy, D.R. Lovley, L.M. Tender, Cyclic voltammetryof biofilms of wild type and mutant Geobacter sulfurreducens on fuel cell anodes in-dicates possible roles of OmcB, OmcZ, type IV pili, and protons in extracellular elec-tron transfer, Energy Environ. Sci. 2 (2009) 506–516.

[19] D.R. Lovley, The microbe electric: conversion of organic matter to electricity, Curr.Opin. Biotechnol. 19 (2008) 564–571.

[20] D. Millo, F. Harnisch, S.A. Patil, H.K. Ly, U. Schröder, P. Hildebrandt, In situ spec-troelectrochemical investigation of electrocatalytic microbial biofilms bysurface-enhanced resonance Raman spectroscopy, Angew. Chem. Int. Ed. 50 (2011)2625–2627.

[21] S.M. Strycharz, A.P. Malanoski, R.M. Snider, H. Yi, D.R. Lovley, L.M. Tender, Appli-cation of cyclic voltammetry to investigate enhanced catalytic current generation


by biofilm-modified anodes of Geobacter sulfurreducens strain DL1 vs. variantstrain KN400, Energy Environ. Sci. 4 (2011) 896–913.

[22] K. Inoue, C. Leang, A.E. Franks, T.L. Woodard, K.P. Nevin, D.R. Lovley, Specific local-ization of the c-type cytochrome OmcZ at the anode surface in current-producingbiofilms of Geobacter sulfurreducens, Environ. Microbiol. Rep. 3 (2011) 211–217.

[23] G. Reguera, K.D. McCarthy, T. Mehta, J.S. Nicoll, M.T. Tuominen, D.R. Lovley, Extra-cellular electron transfer via microbial nanowires, Nature 435 (2005) 1098–1101.

[24] M.Y. El-Naggar, G. Wanger, K.M. Leung, T.D. Yuzvinsky, G. Southam, J. Yang, W.M.Lau, K.H. Nealson, Y.A. Gorby, Electrical transport along bacterial nanowires fromShewanella oneidensisMR-1, Proc. Natl. Acad. Sci. U. S. A. 107 (2010) 18127–18131.

[25] Y.A. Gorby, S. Yanina, J.S. McLean, K.M. Rosso, D. Moyles, A. Dohnalkova, T.J.Beveridge, I.S. Chang, B.H. Kim, K.S. Kim, D.E. Culley, S.B. Reed, M.F. Romine,D.A. Saffarini, E.A. Hill, L. Shi, D.A. Elias, D.W. Kennedy, G. Pinchuk, K. Watanabe,S.i. Ishii, B. Logan, K.H. Nealson, J.K. Fredrickson, Electrically conductive bacterialnanowires produced by Shewanella oneidensis strain MR-1 and other microorgan-isms, Proc. Natl. Acad. Sci. U. S. A. 103 (2006) 11358–11363.

[26] X. Jiang, J. Hu, L.A. Fitzgerald, J.C. Biffinger, P. Xie, B.R. Ringeisen, C.M. Lieber, Probingelectron transfer mechanisms in Shewanella oneidensis MR-1 using a nanoelectrodeplatformand single-cell imaging, Proc. Natl. Acad. Sci. U. S. A. 107 (2010) 16806–16810.

[27] E. Marsili, D.B. Baron, I.D. Shikhare, D. Coursolle, J.A. Gralnick, D.R. Bond,Shewanella secretes flavins that mediate extracellular electron transfer, Proc.Natl. Acad. Sci. U. S. A. 105 (2008) 3968–3973.

[28] L.A. Meitl, C.M. Eggleston, P.J.S. Colberg, N. Khare, C.L. Reardon, L. Shi, Electro-chemical interaction of Shewanella oneidensisMR-1 and its outer membrane cyto-chromes OmcA and MtrC with hematite electrodes, Geochim. Cosmochim. Acta73 (2009) 5292–5307.

[29] D. Baron, E. LaBelle, D. Coursolle, J.A. Gralnick, D.R. Bond, Electrochemical mea-surement of electron transfer kinetics by Shewanella oneidensis MR-1, J. Biol.Chem. 284 (2009) 28865–28873.

[30] M. Sun, F. Zhang, Z.-H. Tong, G.-P. Sheng, Y.-Z. Chen, Y. Zhao, Y.-P. Chen, S.-Y.Zhou, G. Liu, Y.-C. Tian, H.-Q. Yu, A gold-sputtered carbon paper as an anode forimproved electricity generation from a microbial fuel cell inoculated withShewanella oneidensis MR-1, Biosens. Bioelectron. 26 (2010) 338–343.

[31] A. Okamoto, R. Nakamura, K. Hashimoto, In-vivo identification of direct electrontransfer from Shewanella oneidensis MR-1 to electrodes via outer-membraneOmcA-MtrCAB protein complexes, Electrochim. Acta 56 (2011) 5526–5531.

[32] O. Bretschger, A.C.M. Cheung, F. Mansfeld, K.H. Nealson, Comparative microbialfuel cell evaluations of Shewanella spp. Electroanalysis 22 (2010) 883–894.

[33] H.J. Kim, H.S. Park, M.S. Hyun, I.S. Chang, M. Kim, B.H. Kim, A mediator-less micro-bial fuel cell using a metal reducing bacterium, Shewanella putrefaciens, EnzymeMicrob. Technol. 30 (2002) 145–152.

[34] H. Kim, M. Hyun, I. Chang, B. Kim, A microbial fuel cell type lactate biosensorusing a metal—reducing bacterium, Shewanella putrefaciens, J. Microbiol. Bio-technol. 9 (1999) 365–367.

[35] B.H. Kim, H.J. Kim, M.S. Hyun, D.H. Park, Direct electrode reaction of Fe(III)-reducingbacterium, Shewanella putrefaciens, J. Microbiol. Biotechnol. 9 (1999) 127–131.

[36] R. Nakamura, F. Kai, A. Okamoto, G.J. Newton, K. Hashimoto, Self-constructed electri-cally conductive bacterial networks, Angew. Chem. Int. Ed. 48 (2009) 508–511.

[37] A. Okamoto, R. Nakamura, K. Ishii, K. Hashimoto, In vivo electrochemistry of c-typecytochrome-mediated electron-transfer with chemical marking, ChemBioChem 10(2009) 2329–2332.

[38] R. Nakamura, K. Ishii, K. Hashimoto, Electronic absorption spectra and redoxproperties of C type cytochromes in living microbes, Angew. Chem. Int. Ed. 48(2009) 1606–1608.

[39] Y. Zhao, K. Watanabe, R. Nakamura, S. Mori, H. Liu, K. Ishii, K. Hashimoto,Three-dimensional conductive nanowire networks for maximizing anode perfor-mance in microbial fuel cells, Chem. Eur. J. 16 (2010) 4982–4985.

[40] S.-L. Li, S. Freguia, S.-M. Liu, S.-S. Cheng, S. Tsujimura, O. Shirai, K. Kano, Effects ofoxygen on Shewanella decolorationis NTOU1 electron transfer to carbon-felt elec-trodes, Biosens. Bioelectron. 25 (2010) 2651–2656.

[41] X. Li, Y. Li, F. Li, S. Zhou, C. Feng, T. Liu, Interactively interfacial reaction ofiron-reducing bacterium and goethite for reductive dechlorination of chlorinatedorganic compounds, Chin. Sci. Bull. 54 (2009) 2800–2804.

[42] J.C. Biffinger, L.A. Fitzgerald, R. Ray, B.J. Little, S.E. Lizewski, E.R. Petersen, B.R.Ringeisen, W.C. Sanders, P.E. Sheehan, J.J. Pietron, J.W. Baldwin, L.J. Nadeau, G.R.Johnson, M. Ribbens, S.E. Finkel, K.H. Nealson, The utility of Shewanella japonicafor microbial fuel cells, Bioresour. Technol. 102 (2010) 290–297.

[43] K.L. Turner, M.K. Doherty, H.A. Heering, F.A. Armstrong, G.A. Reid, S.K. Chapman,Redox properties of flavocytochrome c3 from Shewanella frigidimarina NCIMB400,Biochemistry 38 (1999) 3302–3309.

[44] K.L. Pankhurst, C.G. Mowat, E.L. Rothery, J.M. Hudson, A.K. Jones, C.S. Miles, M.D.Walkinshaw, F.A. Armstrong, G.A. Reid, S.K. Chapman, A proton delivery pathwayin the soluble fumarate reductase from Shewanella frigidimarina, J. Biol. Chem.281 (2006) 20589–20597.

[45] J. Huang, B. Sun, X. Zhang, Electricity generation at high ionic strength in micro-bial fuel cell by a newly isolated Shewanella marisflavi EP1, Appl. Microbiol. Bio-technol. 85 (2010) 1141–1149.

[46] C.M. Eggleston, J. Vörös, L. Shi, B.H. Lower, T.C. Droubay, P.J.S. Colberg, Binding anddirect electrochemistry of OmcA, an outer-membrane cytochrome from an iron re-ducing bacterium, with oxide electrodes: a candidate biofuel cell system, Inorg.Chim. Acta 361 (2008) 769–777.

[47] M. Firer-Sherwood, G.S. Pulcu, S.J. Elliott, Electrochemical interrogations of theMtr cytochromes from Shewanella: opening a potential window, J. Biol. Inorg.Chem. 13 (2008) 849–854.

[48] J.K. Fredrickson, M.F. Romine, A.S. Beliaev, J.M. Auchtung, M.E. Driscoll, T.S.Gardner, K.H. Nealson, A.L. Osterman, G. Pinchuk, J.L. Reed, D.A. Rodionov, J.L.M.


http://dx.doi.org/



Rodrigues, D.A. Saffarini, M.H. Serres, A.M. Spormann, I.B. Zhulin, J.M. Tiedje, To-wards environmental systems biology of Shewanella, Nat. Rev. Microbiol. 6(2008) 592–603.

[49] R.S. Hartshorne, B.N. Jepson, T.A. Clarke, S.J. Field, J. Fredrickson, J.M. Zachara, L. Shi,J.N. Butt, D.J. Richardson, Characterization of Shewanella oneidensis MtrC: acell-surface decaheme cytochrome involved in respiratory electron transport to ex-tracellular electron acceptors, J. Biol. Inorg. Chem. 12 (2007) 1083–1094.

[50] Y. Xiong, L. Shi, B. Chen, M.U. Mayer, B.H. Lower, Y. Londer, S. Bose, M.F. Hochella,J.K. Fredrickson, T.C. Squier, High-affinity binding and direct electron transfer tosolid metals by the Shewanella oneidensis MR-1 outer membrane c-type cyto-chrome OmcA, J. Am. Chem. Soc. 128 (2006) 13978–13979.

[51] A.S. Beliaev, D.M. Klingeman, J.A. Klappenbach, L. Wu, M.F. Romine, J.M. Tiedje,K.H. Nealson, J.K. Fredrickson, J. Zhou, Global transcriptome analysis of Shewanellaoneidensis MR-1 exposed to different terminal electron acceptors, J. Bacteriol. 187(2005) 7138–7145.

[52] Hong, G. Yi, G.U.O. Jun, C. Xu, Y. Zhi, Sun Mei, P. Guo, Humic substances act aselectron acceptor and redox mediator for microbial dissimilatory azoreductionby Shewanella decolorationis S12, J. Microbiol. Biotechnol. (2007) 428–437.

[53] C.E. Turick, L.S. Tisa, J.F. Caccavo, Melanin production and use as a soluble electronshuttle for Fe(III) oxide reduction and as a terminal electron acceptor byShewanella algae BrY, Appl. Environ. Microbiol. 68 (2002) 2436–2444.

[54] C.E. Turick, A.S. Beliaev, B.A. Zakrajsek, C.L. Reardon, D.A. Lowy, T.E. Poppy, A.Maloney,A.A. Ekechukwu, The role of 4-hydroxyphenylpyruvate dioxygenase in enhancementof solid-phase electron transfer by Shewanella oneidensisMR-1, FEMS Microbiol. Ecol.68 (2009) 223–225.

[55] D.P. Lies, M.E. Hernandez, A. Kappler, R.E. Mielke, J.A. Gralnick, D.K. Newman,Shewanella oneidensis MR-1 uses overlapping pathways for iron reduction at adistance and by direct contact under conditions relevant for biofilms, Appl. Envi-ron. Microbiol. 71 (2005) 4414–4426.

[56] H. von Canstein, J. Ogawa, S. Shimizu, J.R. Lloyd, Secretion of flavins by Shewanellaspecies and their role in extracellular electron transfer, Appl. Environ. Microbiol.74 (2008) 615–623.

[57] S.B. Velasquez-Orta, I.M. Head, T.P. Curtis, K. Scott, J.R. Lloyd, H. Von Canstein, Theeffect of flavin electron shuttles in microbial fuel cells current production, Appl.Environ. Microbiol. 85 (2010) 1373–1381.

[58] R.A. Bouhenni, G.J. Vora, J.C. Biffinger, S. Shirodkar, K. Brockman, R. Ray, P. Wu,B.J. Johnson, E.M. Biddle, M.J. Marshall, L.A. Fitzgerald, B.J. Little, J.K. Fredrickson,A.S. Beliaev, B.R. Ringeisen, D.A. Saffarini, The role of Shewanella oneidensis MR-1outer surface structures in extracellular electron transfer, Electroanalysis 22(2010) 856–864.

[59] Y. Jiao, F. Qian, Y. Li, G. Wang, C.W. Saltikov, J.A. Gralnick, Deciphering the electrontransport pathway for graphene oxide reduction by Shewanella oneidensis MR-1,J. Bacteriol. 193 (2011) 3662–3665.

[60] G.J. Newton, S. Mori, R. Nakamura, K. Hashimoto, K. Watanabe, Analyses ofcurrent-generating mechanisms of Shewanella loihica PV-4 and Shewanellaoneidensis MR-1 in microbial fuel cells, Appl. Environ. Microbiol. 75 (2009)7674–7681.

[61] F. Harnisch, K. Rabaey, The diversity of techniques to study electrochemicallyactive biofilms highlights the need for standardization, ChemSusChem. specialissue "Microbial fuel cell" (in press), http://dx.doi.org/10.1002/cssc.201100817.

[62] C.I. Torres, R. Krajmalnik-Brown, P. Parameswaran, A.K.Marcus, G.Wanger, Y.A. Gorby,B.E. Rittmann, Selecting anode-respiring bacteria based on anode potential: phyloge-netic, electrochemical, and microscopic characterization, Energy Environ. Sci. 43(2009) 9519–9524.

[63] E.J. Cho, A.D. Ellington, Optimization of the biological component of abioelectrochemical cell, Bioelectrochemistry 70 (2007) 165–172.

[64] A.A. Carmona-Martínez, F. Harnisch, L.A. Fitzgerald, J.C. Biffinger, B.R. Ringeisen,U. Schröder, Cyclic voltammetric analysis of the electron transfer of Shewanellaoneidensis MR-1 and nanofilament and cytochrome knock-out mutants, Bio-electrochemistry 81 (2011) 74–80.

[65] H.W. Harris, M.Y. El-Naggar, O. Bretschger, M.J. Ward, M.F. Romine, A.Y. Obraztsova,K.H. Nealson, Electrokinesis is a microbial behavior that requires extracellular elec-tron transport, Proc. Natl. Acad. Sci. U. S. A. 107 (2010) 326–331.

[66] H. Liu, S. Matsuda, S. Kato, K. Hashimoto, S. Nakanishi, Redox-responsiveswitching in bacterial respiratory pathways involving extracellular electrontransfer, ChemSusChem 3 (2010) 1253–1256.

[67] H. Liu, S. Matsuda, T. Kawai, K. Hashimoto, S. Nakanishi, Feedback stabilization in-volving redox states of c-type cytochromes in living bacteria, Chem. Commun. 47(2011) 3870–3872.

[68] J.T. Babauta, H.D. Nguyen, H. Beyenal, Redox and pH microenvironments withinShewanella oneidensis MR-1 biofilms reveal an electron transfer mechanism,Energy Environ. Sci. 45 (2011) 6654–6660.

[69] B.R. Ringeisen, E. Henderson, P.K. Wu, J. Pietron, R. Ray, B. Little, J.C. Biffinger, J.M.Jones-Meehan, High power density from a miniature microbial fuel cell usingShewanella oneidensis DSP10, Energy Environ. Sci. 40 (2006) 2629–2634.


[70] O. Bretschger, A. Obraztsova, C.A. Sturm, S.C. In, Y.A. Gorby, S.B. Reed, D.E. Culley,C.L. Reardon, S. Barua, M.F. Romine, J. Zhou, A.S. Beliaev, R. Bouhenni, D. Saffarini,F. Mansfeld, B.H. Kim, J.K. Fredrickson, K.H. Nealson, Current production andmetal oxide reduction by Shewanella oneidensis MR-1 wild type and mutants,Appl. Environ. Microbiol. 73 (2007) 7003–7012.

[71] R.M. Atlas, Handbook of Microbiological Media, third ed. CRC Press LLC, Florida,1993.

[72] D. Coursolle, D.B. Baron, D.R. Bond, J.A. Gralnick, The Mtr respiratory pathway isessential for reducing flavins and electrodes in Shewanella oneidensis, J. Bacteriol.(2010) (JB.00925-00909).

[73] H. Pivnick, Pseudomonas rubescens, a new species from soluble oil emulsions,J. Bacteriol. 70 (1955) 1–6.

[74] K.H. Nealson, C.R. Myers, Microbial reduction of manganese and iron: new ap-proaches to carbon cycling, Appl. Environ. Microbiol. 58 (1992) 439–443.

[75] B.F. Vogel, K. Jørgensen, H. Christensen, J.E. Olsen, L. Gram, Differentiation ofShewanella putrefaciens and Shewanella alga on the basis of whole-cell proteinprofiles, ribotyping, phenotypic characterization, and 16S rRNA gene sequenceanalysis, Appl. Environ. Microbiol. 63 (1997) 2189–2199.

[76] K. Venkateswaran, D.P. Moser, M.E. Dollhopf, D.P. Lies, D.A. Saffarini, B.J.MacGregor, D.B. Ringelberg, D.C. White, M. Nishijima, H. Sano, J. Burghardt, E.Stackebrandt, K.H. Nealson, Polyphasic taxonomy of the genus Shewanella anddescription of Shewanella oneidensis sp. nov, Int. J. Syst. Bacteriol. 49 (1999)705–724.

[77] R.J. Owen, R.M. Legros, S.P. Lapage, Base composition, size and sequence similar-ities of genome deoxyribonucleic acids from clinical isolates of Pseudomonasputrefaciens, J. Gen. Microbiol. 104 (1978) 127–138.

[78] J.C. Biffinger, R. Ray, B.J. Little, L.A. Fitzgerald, M. Ribbens, S.E. Finkel, B.R.Ringeisen, Simultaneous analysis of physiological and electrical output changesin an operating microbial fuel cell with Shewanella oneidensis, Biotechnol. Bioeng.103 (2009) 524–531.

[79] M. Rosenbaum, M.A. Cotta, L.T. Angenent, Aerated Shewanella oneidensis in con-tinuously fed bioelectrochemical systems for power and hydrogen production,Biotechnol. Bioeng. 105 (2010) 880–888.

[80] V. Fourmond, K.Hoke, H.A. Heering, C. Baffert, F. Leroux, P. Bertrand, C. Léger, SOAS: afree program to analyze electrochemical data and other one-dimensional signals,Bioelectrochemistry 76 (2009) 141–147.

[81] T.R. Neu, B. Manz, F. Volke, J.J. Dynes, A.P. Hitchcock, J.R. Lawrence, Advanced im-aging techniques for assessment of structure, composition and function in biofilmsystems, FEMS Microbiol. Ecol. 72 (2010) 1–21.

[82] C. Staudt, H. Horn, D.C. Hempel, T.R. Neu, Volumetric measurements of bacterialcells and extracellular polymeric substance glycoconjugates in biofilms, Bio-technol. Bioeng. 88 (2004) 585–592.

[83] Y. Yang, G. Sun, J. Guo, M. Xu, Differential biofilms characteristics of Shewanelladecolorationis microbial fuel cells under open and closed circuit conditions, Bio-resour. Technol. 102 (2011) 7093–7098.

[84] K. Fricke, F. Harnisch, U. Schroder, On the use of cyclic voltammetry for the studyof anodic electron transfer in microbial fuel cells, Energy Environ. Sci. 1 (2008)144–147.

[85] E. Marsili, J.B. Rollefson, D.B. Baron, R.M. Hozalski, D.R. Bond, Microbial biofilmvoltammetry: direct electrochemical characterization of catalytic electrode-attachedbiofilms, Appl. Environ. Microbiol. 74 (2008) 7329–7337.

[86] L. Shi, B. Chen, Z. Wang, D.A. Elias, M.U. Mayer, Y.A. Gorby, S. Ni, B.H. Lower, D.W.Kennedy, D.S. Wunschel, H.M. Mottaz, M.J. Marshall, E.A. Hill, A.S. Beliaev, J.M.Zachara, J.K. Fredrickson, T.C. Squier, Isolation of a high-affinity functional proteincomplex between OmcA and MtrC: two outer membrane decaheme c-type cyto-chromes of Shewanella oneidensis MR-1, J. Bacteriol. 188 (2006) 4705–4714.

[87] J. Wang, M. Li, Z. Shi, N. Li, Z. Gu, Direct electrochemistry of cytochrome c at aglassy carbon electrode modified with single-wall carbon nanotubes, Anal.Chem. 74 (2002) 1993–1997.

[88] L. Wang, E. Wang, Direct electron transfer between cytochrome c and a goldnanoparticles modified electrode, Electrochem. Commun. 6 (2004) 49–54.

[89] T.L. Miller, M.J. Wolin, A serum bottle modification of the Hungate technique forcultivating obligate anaerobes, J. Appl. Microbiol. 27 (1974) 985–987.

[90] A.E. Franks, K.P. Nevin, H. Jia, M. Izallalen, T.L. Woodard, D.R. Lovley, Novel strat-egy for three-dimensional real-time imaging of microbial fuel cell communities:monitoring the inhibitory effects of proton accumulation within the anode bio-film, Energy Environ. Sci. 2 (2009) 113–119.

[91] Z. Lewandowski, H. Beyenal, D. Stookey, Reproducibility of biofilm processes andthe meaning of steady state in biofilm reactors, Water Sci. Technol. (2004)359–364.

[92] K.H. Williams, K.P. Nevin, A. Franks, A. Englert, P.E. Long, D.R. Lovley, Electro-de-based approach for monitoring in situ microbial activity during subsurfacebioremediation, Energy Environ. Sci. 44 (2009) 47–54.



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Electron transfer and biofilm formation of Shewanella putrefaciens as function of anode potential