Interfacial electron transfer between Geobacter sulfurreducens and gold electrodes via carboxylate-alkanethiol linkers: Effects of the linker length

Bioelectrochemistry 126 (2019) 130–136

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Interfacial electron transfer between Geobacter sulfurreducens and gold electrodes via carboxylate-alkanethiol linkers: Effects of the linker length M. Füeg a,⁎,1, Z. Borjas b, M. Estevez-Canales c, A. Esteve-Núñez b,c, I.V. Pobelov a, P. Broekmann a, A. Kuzume a,⁎,2 a b c

Department of Chemistry and Biochemistry, University of Bern, Freiestrasse 3, Bern 3012, Switzerland IMDEA WATER, Alcalá de Henares, Madrid, Spain Department of Chemical Engineering, University of Alcalá, Alcalá de Henares, Madrid, Spain

a r t i c l e

i n f o

Article history: Received 14 May 2018 Received in revised form 29 November 2018 Accepted 30 November 2018 Available online 12 December 2018 Keywords: Geobacter sulfurreducens SAM Microbial fuel cells Extracellular electron transfer Outer membrane cytochromes

a b s t r a c t Geobacter sulfurreducens (Gs) attachment and biofilm formation on self-assembled monolayers (SAMs) of carboxyl-terminated alkanethiol linkers with varied chain length on gold (Au) was investigated by electrochemical and microscopic methods to elucidate the effect of the surface modification on the current production efficiency of Gs cells and biofilms. At the initial stage of the cell attachment, the electrochemical activity of Gs cells at a submonolayer coverage on the SAM-Au surface was independent of the linker length. Subsequently, multiple potential cyclings indicated that longer linkers provided more biocompatible conditions for Gs cells than shorter ones. For Gs biofilms, on the other hand, the turnover current decreased exponentially with the linker length. During the biofilm formation, bacteria need to adjust from the initial planktonic state to an electrode-respiring state, which was triggered by a strong electrochemical stress found for shorter linkers, resulting in the formation of mature biofilms. Our results suggest that the initial cell attachment and the biofilm formation are two inherently different processes. Therefore, the effects of linker molecules, electron transfer efficiency and biocompatibility, must be explored simultaneously to understand both processes to increase the current production of electrogenic microorganisms in microbial fuel cells. © 2018 Elsevier B.V. All rights reserved.

1. Introduction The discovery of electroactive bacteria has been fundamental to the research of the electrochemical interface between bacteria and electro-conductive materials, and led to the invention of Microbial Electrochemical Technologies (MET) [1]. A large variety of applications have been developed on this basis: direct power generation (microbial fuel cells, MFCs) [2,3]; chemical production of H2 (microbial electrolysis cells, MECs) [4,5]; microbial electrosynthesis [6,7]; biocathodes-based nutrient removal such as nitrate [7]; water desalination (microbial desalination cells, MDCs) [8,9]; bioelectrochemical constructed wetlands (METlands) [10] and microbial electroremediating cells (MERC) for restoring polluted environments [11,12]. Several parameters of MET, such as reactor configuration, species of bacteria and electrolyte composition, have been optimized to maximize performance of microbial electrochemical devices [13–15]. In this same context, the fundamental understanding of the interfacial processes between bacteria and

⁎ Corresponding authors. E-mail addresses: [email protected] (M. Füeg), [email protected] (A. Kuzume). 1 Current affiliation: Department of Chemical Engineering, University of Alcalá. 2 Current affiliation: Institute of Innovative Research, Tokyo Institute of Technology.

https://doi.org/10.1016/j.bioelechem.2018.11.013 1567-5394/© 2018 Elsevier B.V. All rights reserved.

electrodes is essential. Therefore, it is important to establish an efficient “electric communication” between the electroactive bacteria and the anode, thereby lowering the resistance of the interface and leading to an overall higher microbial current production. The bacteria of genus Geobacter sulfurreducens (Gs), which produces the highest current densities of all known pure cultures, represents the most thoroughly investigated family of electroactive bacteria [16–18]. In particular, the short-range electron transfer (ET) between individual cells of Gs and an electron-accepting electrode (anode) is proposed to be dominated by the outer membrane c-type cytochromes (OMCs) [19]. Several studies have shown that OMCs on the Gs surface are responsible for the direct ET to metal electrodes [20–24], a process that fails when the amount of OMCs is strongly reduced [25]. Among them, OmcZ is supposed to be a key OMC for a direct ET to metal electrodes [19,20]. Indications of mediator-less electron transfer were shown by other authors [25–27]. Previously, we reported on the electrochemical activity and structural properties of Gs cells directly attached on bare gold electrodes using a combination of in situ spectroscopic, microscopic and electrochemical methods [22]. We presented clear evidences that OMCs change their structure and electrochemical properties as a function of the electrode potential when bacteria are in close contact with Au surfaces [22]. Furthermore, when Au surfaces were modified with self-assembled monolayers (SAMs) of ω-functionalized

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decanethiols containing different outer linking groups, the interfacial electrochemical and spectroscopic studies revealed that carboxylterminated alkanethiols enhanced microbial current production [23]. In the current paper, we used electrochemical and microscopic techniques to investigate the effect of the length of the carboxyl-terminated alkanethiols on (i) the initial attachment process of Gs on SAM-modified Au, (ii) the further growing process of Gs biofilms and (iii) the interfacial electron transfer between Gs and Au. 2. Experimental 2.1. Chemicals and materials Horse heart cytochrome c (cyt c), 2-mercaptoacetic acid (C1), 3-mercaptopropionic acid (C2), 6-mercaptohexanoic acid (C5), 8-mercaptooctanoic acid (C7), 11-mercaptoundecanoic acid (C10), 16-mercaptohexadecanoic acid (C15), 70% glutaraldehyde, NaHCO3, NH4Cl, KCl and ethanol (analytical grade) were purchased from Sigma Aldrich. MilliQ water (Total Organic Carbon value: TOC b 4 ppb, 18.2 MΩcm) was used throughout the work. N2 + CO2 gas mixture (Carbagas: 80:20 (v:v)) was used to maintain the electrolyte anaerobically at pH 6.8. The gas was filtered through a Varian CP17970 filter to eliminate oxygen traces before entering the electrochemical cell. Gs was anaerobically cultured in a serum bottles containing a freshwater medium (2.5 g/L NaHCO3, 0.36 g/L NaH2PO4, 0.50 g/L NH4Cl and 0.10 g/L KCl) supplemented with minerals and vitamins as described elsewhere [22,23]. Growth was supported by 20 mM sodium acetate as the sole carbon source and electron donor, and 40 mM sodium fumarate serving as a terminal electron acceptor [23]. Furthermore, 0.1 mM Fe(II) was supplied in excess to avoid limiting the synthesis of cytochromes by insufficient Fe(II) supply. The serum bottles were sterilized and purged with a N2:CO2 (80:20 (v:v)) gas mixture to keep the pH of the media constant at 6.8. For electroactive biofilm growth experiments, the same freshwater medium was used. Growth was supported by 20 mM sodium acetate as the sole carbon source and electron donor, while an Au electrode was used as a terminal electron acceptor instead of fumarate. For cyt c studies, an argon-degassed 5 mM phosphate buffer (containing 0.10 g/L NaCl, 0.35 g/L NaH2PO4 and 0.36 g/L Na2HPO4) was used. Degassing was maintained during the assays. All employed electrodes were polycrystalline Au Clavillier-type [28] beads (cyt c experiments), half-beads (Gs biofilm) or disk electrodes (Gs sub-monolayer (subML)). The half-bead and bead electrodes were cleaned by three rounds of electrochemical polishing [29], followed by subsequent flame annealing for 30 s to dark-red glow and cooled down to room temperature before each experiment. The disk electrodes were polished with 1 μm polycrystalline diamond crystals (MetaDi, Buehler) and electropolished [29] to obtain clean surfaces. SAMs were formed by immersing the Au surfaces in 1 mM solution of the corresponding linker in ethanol overnight at 65 °C, then rinsing in ethanol to remove loosely bound linker molecules, and finally drying in a stream of argon before mounting in the electrochemical cell. For the cyt c assembly, the SAM-modified electrode was immersed into a 50 μM cyt c in 5 mM phosphate buffer for 1 h at 4 °C.

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All glassware and Teflon parts were cleaned by boiling in 25% HNO3 for 30 min and subsequently boiled three times in MilliQ water, followed by excessive rinsing with MilliQ water between each boiling step. The electrolytes and parts of the electrochemical setup (except the reference and working electrodes) used for the biofilm experiments were sterilized at 125 °C and 1.40 bar for 20 min and cooled to 33 °C prior to use. The assembly of the electrochemical cell and injection of the Gs solution have been conducted next to a Bunsen burner under a vented hood to avoid contamination. Experiments with cyt c where performed with an Autolab PGSTAT 30 Potentiostat (Metrohm Autolab, Netherlands) in a lab-made three compartment electrochemical cell; the solution resistance was compensated. The current densities were calculated using either the geometric (disk electrodes) or the electrochemically determined (bead electrodes, half-bead electrodes) surface areas. 2.3. SubML assembly and biofilm growth 5 mL of Gs solution (optical density OD of 0.5, corresponding to a cell density of 0.7 pM or cell number of 4 × 108 cells/mL) were added to 170 mL of freshwater medium containing 20 mM sodium acetate and purged with a 80:20 (v:v) N2 + CO2 gas mixture to remove oxygen and maintain solution pH at 6.8. Gs cells were then attached on linker-modified electrodes by polarizing at 0.20 V for 16 h. During the polarization, the solution was stirred with a magnetic stir bar. After 16 h of polarization, the subML of Gs was examined with 15 subsequent potential cycles between −0.65 and 0.20 V at a scan rate of 0.01 V s−1. For growing the electroactive biofilm, the electrochemical cell was continuously fed with sterile medium containing 3.6 μM Fe(II) from a 2 L tank. Both the anoxic conditions and the pH (6.8) of the inlet media were kept constant by purging with the 80:20 (v:v) N2 + CO2 gas mixture through a Minisart NML Plus filter (0.2 μm pore size, Sartorius). A peristaltic pump (Longerpump BT100-2 J) ensured a continuous supply of the growth media with a solution exchange rate of 0.021 h−1. The growth media was supplied only after reaching a current density of 8 μA cm−2. During the growth of the biofilms in the electrochemical cell, the potential was fixed at 0.20 V, the solution was gently stirred, and the headspace was continuously purged with the N2 + CO2 gas mixture. Every submonolayer and biofilm experiment was repeated a minimum three times, and the average values for current production were compared to confirm the reproducibility of the experiments. 2.4. Scanning electron microscopy (SEM) The coverage of the microbial layer was examined with a HITACHI S-3000 N Scanning Electron Microscope operated at 25 kV. Cellcolonized electrodes were first immersed in a 2% (v/v) glutaraldehyde solution for 30 min, subsequently desalted in a water:ethanol dilution series (100:0, 75:25, 50:50, 25:75, 0:100) for 10 min each, and dried in air (adapted from Ref. [18]). The samples were sputter coated with a 4 nm Au film before mounting into the SEM chamber. The relative coverage of Gs cell was calculated assuming a surface occupation area of 0.96 μm2 per bacterial cell [23].

2.2. Electrochemical experiments

3. Results

All electrochemical experiments with Gs were conducted in labmade electrochemical cells that accommodated eight working electrodes, an Au-wire counter electrode and a Ag/AgCl (sat. KCl) reference electrode. The cell was fitted with inlets and outlets for gas and solution exchange. All potentials are quoted against this reference. A CHI1030c multi-potentiostat (CH Instruments, USA) was used for electrochemical measurements. The temperature was maintained at 33 °C throughout all Gs experiments. Cyt c experiments were performed at room temperature.

3.1. In vivo electrochemical response of submonolayers of bacteria linked to Au To investigate the initial stage of biofilm formation, a subML coverage of Gs was prepared by polarizing a bare Au and Cn-Au surfaces at 0.20 V for 16 h in a medium as described in section 2.1. The resulting current transients were mostly similar to the ones previously reported [22,23], i.e. after inoculation of Gs (20 pM final concentration), a steep increase of the current was followed by a slow decrease over hours

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(Fig. S2). SEM images observed after 16 h of polarization showed a coverage of 8–10 cells per 100 μm2 regardless of the Cn-Au surface used (Fig. S3 upper panel and Table S1). To further explore the electrochemical activity of Gs subML on the Cn-Au surfaces, a set of 15 subsequent CVs was recorded after colonization (Fig. S4). Fig. 1A shows the first CVs of Gs subML attached on the Cn-Au surfaces, representing their electrochemical activity. They showed no differences or overlaps for all linkers except the longest one (n = 15). The formal redox potential of Gs on Cn-Au surfaces (1 ≤ n ≤ 10) was −0.30 to −0.31 V (Table S2), which is in the range of the formal redox potentials of OMCs for Gs biofilms, as reported in previous electrochemical and spectroscopic studies [20,22,23,30−32]. Consequently, we attributed the observed CV peaks to the reduction and re-oxidation of the OMCs. Interestingly, when Gs subML were subjected to 15 potential cycles of CVs (Fig. 1B), the resulting voltammetric features vary significantly with the linker length. For all surfaces, the total charge of the reduction peaks decreased after 15 cycles below 5% of the value from the 1st cycle (Fig. S5). In addition, the formal redox potentials shifted by 0.14–0.31 V toward the positive potential range (Fig. S6C and Table S2). SEM images of Gs subML were recorded after 15 potential cycles, for all surface types, to visualize the effect of potential cycling on the coverage of Gs cells. However, no significant changes were detected (Figs. 1D and S3 lower panel, summarized in Table S1). We further noted that the voltammetric features obtained for the first cycle can be recovered for all surfaces after 15 potential cycles by repolarizing at 0.20 V for 1 h, as also previously reported for a Gs subML formed on a bare Au anode [22]. 3.2. In vivo electrochemical response of bacteria biofilm linked to Au The quasi-steady state CVs of six-days old Gs biofilms on both bare and Cn-Au surfaces reveal a clear turnover feature associated with the acetate metabolism (Fig. 2); CVs for all linkers except C15 reaching saturation at potentials below 0.30 V. The half-saturation potential [33] varied with the linker length: i.e. for n = 0–7, it was located between −0.36 and − 0.33 V; in contrast, for n = 10 the potential was

Fig. 2. Representative cyclic voltammograms of Gs biofilms as a function of chain length. Scan rate: 0.001 V s−1.

−0.27 V (Fig. S7A and Table S3). The limiting current is a measure for the rate of the acetate oxidation by Gs, the values measured at 0.30 V (average of 3 independent experiments) were shown to decrease with the increase of the chain length (Fig. 3). For comparison, Fig. 3 also includes the maximum current measured on the C15-Au surface at E = 0.30 V. For better illustration of this trend, the reader is referred to Fig. S7B. We point out here that according to Fig. 2, the C7-Au surface shows a higher limiting current compared to the C5-Au surface, which is contradictory to the trend shown in Fig. 3. This is due to a low experimental reproducibility originating from living samples. Therefore, all electrochemical experiments were repeated three times and the average limiting current values were plotted in Fig. 3, where the inconsistencies are represented by the large error bars. The SEM image of a six-days old Gs biofilm on a C1-Au surface polarized at 0.20 V showed a dense biofilm (Fig. 3A) in contrast with the Gs adlayer observed on the C5-Au surface (Fig. 3B). This shows minor aggregation of the cells but still has a subML coverage. The SEM image of Gs adsorbed on a C15-Au surface showed individual Gs cells homogeneously distributed with a surface coverage of ca. 11 cells per 100 μm2 (Fig. 3C) which is similar to that of a subML obtained after 16 h of

Fig. 1. Cyclic voltammograms of the Gs SubML on bare Au (dashed line) and Cn-Au (solid lines) surfaces recorded at 0.01 V s−1: the first (A) and the 15th cycle (B). SEM images of the subML formed on C7-Au before (C) and after (D) potential cycling, as obtained in independent experiments.

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Fig. 3. The limiting current obtained at 0.30 V from the CVs of biofilms as a function of chain length. The SEM images (A-C) are taken after performing the CVs on the indicated Cn-Au surfaces. Scale bar: 10 μm.

polarization (Fig. S3). Other SEM images of biofilms formed on Cn-Au surfaces are provided in Fig. S8.

4. Discussion 4.1. Linker length effect on a Gs subML The successful formation of a Gs subML on the Cn-Au surfaces, as the initial step of the biofilm formation, was demonstrated by voltammetric and SEM studies. The overlapping of the first CVs with that on a bare Au electrode (Fig. 1A) surprisingly indicates that the linker's length (1 ≤ n ≤ 10) does not influence the electrochemical properties of OMCs of the attached Gs cells, as well as the accumulation process of Gs cells on the polarized electrode surfaces. From here we exclude the experimental data of C15-Au from further discussion since, judging from a large peak-to-peak separation of the reduction and re-oxidation peaks (Fig. 1A) and results for cyt c (Fig. S1), the ET process is limited by electron tunneling through a thick SAM and is much slower than on the other surfaces.

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Due to the continuous evolution of the system, we were not able to obtain the ET rate constant (kET) values of the Gs subMLs formed on bare Au and Cn-Au surfaces using Laviron's method. However, we still can draw certain conclusions on the kinetics of the OMCs reduction/reoxidation based on the peak-to-peak separation, which was 0.33 V at a scan rate of 0.01 V·s−1 for Gs on Cn-Au (1 ≤ n ≤ 10). For the cyt c model system on C10-Au at the scan rate of 0.10 V·s−1 it was only 0.01 V (Fig. S1B). This large difference suggests that the redox process for OMCs of Gs is much slower than for the model cyt c. A non-optimal initial configuration, as well as the rigidness of the molecular environment, are expected to increase the required reorganization energy associated with the ET process and to heavily decrease the rate of ET between OMCs and Au when compared to the model cyt c system. There are significant changes in the voltammetric features observed during the first 15 potential cycles of the Gs subML (Fig. S4). First, the formal potential shifts toward more positive values and this trend is more pronounced for the shorter linkers (Figs. 4A). Second, the separation between the reduction and re-oxidation peak decreases for all surfaces from 0.33 V to 0.24–0.27 V between the first and second cycle. Subsequently, it increases up to 0.36 V for C1-Au, remains rather constant for bare gold, and decreases for the other Cn-Au surfaces (Fig. 4B). Third, the charge of the peaks corresponding to the reduction and subsequent re-oxidation of OMCs decreases significantly during the cycling (Fig. 4C and D). Both the reduction and re-oxidation charges steadily decrease during each cycle by 65–83% of the previous value. This decrease in charge cannot be attributed to a change in the Gs cell coverage, as demonstrated by SEM studies (Fig. S3 and Table S1). The decrease in charge is more pronounced for short linkers (ca. 65% of the charge is retained per cycle for C1-Au). After 15 cycles, the reduction charge of OMCs on C1-Au is only 0.4% of that in the first cycle, while ca. 5% of charge remains for C7-Au and C10-Au (Fig. S5). We also found that the re-oxidation charge was smaller than the reduction charge in the same cycle, as illustrated for n = 2 in Fig. S9A. This indicates that the electrons transferred to the Gs cells through OMCs during the cathodic scan were partially lost and, therefore, do not contributed to the corresponding anodic processes in the reverse potential scan. Fig. S9B shows the ratio of the reduction and re-oxidation charges Qox/Qred. Only in the cases of bare Au and C1-Au, where the charge ratio

Fig. 4. Detailed analysis of the 15 cycles recorded from the Gs subML formed on the bare and C1-C10 modified Au surfaces. Shown are the formal potential (A), the peak-to-peak separation (B), the logarithm of the reduction (C) and oxidation (D) charge, as function of the cycle number.

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Qox/Qred reaches 1 within 15 cycles, indicates that a steady state is reached. Such a deterioration of the electrochemical activity can be attributed to a “stress” induced on Gs cells by potential cycling. This could be due to such effects as: the change in the electric field strength, protonation/deprotonation of the binding carboxyl group, adsorption/desorption of ions etc. Although the absolute values of the reduction and re-oxidation charges steadily decrease during the potential cycling, an increase of Qox/Qred after 8–10 cycles for all surfaces suggests that, with time, OMCs adapt to this stress. The biophysical background of such an adaptation is most probably due to conformational changes in the protein structure [22,23], which might be reflected as the shift of the formal potential (Fig. 4A). SEM results indicate that the deterioration of charge is not due to the desorption of Gs cells during the potential cycling, therefore we propose that it is caused by changes in the OMC structure, which cannot be detected by SEM. Furthermore, these structural changes appear to be reversible in the long term, because the same CV response as the first CV cycle (shown in Fig. 1A) can be recovered after 15 potential cycles simply by the re-polarization of the anode for one hour. The effect of the SAM formed by linker molecules on Au surfaces can be described as “protection” of Gs cells from the potential cycling stress by decreasing the local electric field or preventing ion adsorption. The packing and local organization of SAMs formed by alkanethiols is mostly driven by intermolecular interactions between alkane chains. Therefore, longer alkanethiols form “more blocking” SAMs of “higher quality” that better protect Gs cells from the electrochemical stresses. In other words, longer linkers provide more biocompatible condition for Gs. The linker length effect on the electrochemical response of Gs subMLs during the potential cycling thus can be explained as following: Gs on bare and Cn-Au (n ≤ 2) surfaces feel a strong stress from potential cycling, therefore OMCs adapt to new conditions quicker (the formal potential is shifting more between potential cycles), but the new structure is not an optimal one (peak-to-peak separation is not decreasing as much, the retained electrochemical activity is very low). Gs subML on SAMs formed by long (n ≥ 5) linkers on the Au surfaces experience a reduced potential cycling stress, and thus their adaptation is slower (the formal potential is shifting less than with the short linkers), but the new structure is a more favorable one (the peak-to-peak separation decreases, the retained electrochemical activity is higher than with the short linkers). Furthermore, we note that all results presented in Figs. 1 and 4 are very similar for C7-Au and C10-Au. We interpret this as an indication that for Gs subML, the protecting effect of the SAM is greater than the reduction of the interfacial ET rate with the increase in linker length (Fig. S1D). 4.2. Linker length effect on the formation and activity of a Gs biofilm The current production by Gs biofilms on the Cn-Au surfaces, as indicated by the acetate-turnover feature, shows a clear dependence on the length of the linker molecules (Fig. 2 and S7). These electrochemical results correlate with SEM findings showing that a thicker biofilm is formed for shorter linkers (Figs. 3 and S8), and for C15-Au, the division of Gs cells seems to be completely suppressed. We explain these results by the dominating role of the kinetics of the interfacial ET between Gs and Au, which can be visualized by normalizing Fig. 2 to the corresponding maximum current (Fig. S7C). In this representation, the curves obtained for bare Au and C1-Au follows the thermodynamic Nernst equation with a half-wave potential at −0.36 V and an effective number of transferred electrons ≈ 0.7. CVs for longer linkers deviate from the Nernstian wave, which can be attributed to a kinetic hindrance. A model for Gs biofilms developed by Strycharz et al. [34] divides the ET from acetate to the electrode into five steps: (1) Acetate uptake into the cytosol; (2) Oxidation of acetate in the cytosol; (3) Chain transfer of the electron to an OMC and its reduction; (4) Electron transport through the biofilm layers; (5) Final electron transfer from the last

OMC to the anode. Cyclic voltammograms for n N 1 in Fig. S7C are similar to model CVs with step 5 as the limiting step, indicating that the interfacial ET between OMCs and the anode through the linker is the ratelimiting step. Although Gs has successfully formed a subML on the Cn-Au surfaces (1 b n ≤ 10), it did not trigger the accumulation of consecutive layers to form a thick biofilm (compare Figs. S3 and S8). We note that the growth of the biofilm is taking place during the polarization at a constant potential, therefore the Gs cells are not permanently affected by the potential cycling stress. We would like to point out the possibility that the variation of the linker length may modify the potential experienced by the cells on SAMs and thus affect the biofilm growth. Our model study using HH cyt-c showed that the redox potential shifted in the positive range with the chain length of linker SAMs (n ≤ 10) in the range of 5–15 mV, similar to those reported by Chidsey et al. for the ferroceneterminated alkanethiol system [35]. Therefore, we cannot exclude the possibility of an effect by potential shifts induced by the different linker lengths, but we do not think that this effect alone would account for the significant difference in the success of biofilm formation. Furthermore, the attenuation of the ET process due to the high resistance of long SAMs can be largely prevented by tuning the electronic properties of the linker molecules. Linker molecules consisting of πconjugated systems, for instance, have a much higher molecular conductance than saturated ones [36]. This may help to overcome the attenuation problem in the ET between microbes and electrode surface, simultaneously providing biocompatible conditions for the microorganisms by utilizing the carboxylic acid group as an anchoring group. It is also important to note that not all MET applications require biofilm growth since certain planktonic cells have been reported to exhibit ET to electrodes. Thus, successful wiring between Gs cells and gold electrodes through conductive linkers constitutes a new scenario for developing biosensor applications where achieving a method for anchoring cells to the electrode is key. In this case, biofilm growth is not a requirement because electroactive cells acting like the “plug and play” cells reported elsewhere [37] could significantly benefit from our linking strategy. 5. Conclusions We investigated the attachment of the electrogenic bacteria Gs on SAMs formed with carboxyl-terminated alkanethiol linkers HS-(CH2) n-COOH with varied chain lengths on gold, and the subsequent biofilm growth. The evolution of the voltammetric features of OMCs in Gs subML (increase in formal potential and peak-to-peak separation, decrease in reduction and re-oxidation charge) during the potential cycling was rationalized as a “stress”. This stress was reduced if long (n ≥ 5) linkers were employed, showing their better biocompatibility. A strong linker effect was found for biofilm formation, where a thicker biofilm with higher electrochemical activity was formed with shorter (n ≤ 2) linkers. Analysis of CVs showing the response to acetate metabolism allowed us to conclude that the ET between Gs in biofilms and the metal electrode was the rate-limiting step. Another factor that might affect biofilm growth is Gs's adaptation from a planktonic state (where fumarate is the terminal electron acceptor) to a surfaceattached state (where electrons are transferred to the electrode), a process accompanied by changes in the expression of OMCs [38,39]. The better biocompatibility of longer linkers might prevent the first layer of Gs cells from “feeling” the surface and activate the transition to the surface-attached state. Our results indicate that the subML and the biofilm of Gs are two inherently different systems with regards to the electrochemical and physiological behavior of the cells on the investigated linker molecules. It is also astonishing that SAMs with the approximate thickness of 0.6–2.4 nm (Table S1) can affect significantly the microbial conglomeration process, which starts with the adherence of 1 μm-sized bacterial cells. This fact highlights the importance of the electrochemical

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environment of the first adherent Gs layer on the electron-accepting electrode surface, which might help to elucidate the mechanism of the biofilm formation of electrogenic microorganisms. We therefore suggest that two effects of linker molecules, ET efficiency and biocompatibility, must be explored simultaneously to improve the attachment condition of Gs cells, and thus to increase the current production of electrogenic microorganisms in microbial fuel cells. Acknowledgements This work was initially started under the supervision of Prof. Thomas Wandlowski (deceased 2015). The support by the European Union through the FP7 BacWire Project (Contract No: MNP4-SL-20092293337), CTI Swiss Competence Centers for Energy Research (SCCER Heat and Electricity Storage), the Swiss National Science Foundation (Grant No. 200020-144471), Swiss Commission for Technology and Innovation (13696.1), COST ActionTD1002, and the University of Bern is acknowledged. MF acknowledges support from an “UniBern Initiator Grant”. The authors are grateful to Prof. Dr. Matthias Arenz (University of Bern) for providing equipment, advice and financial support. AK acknowledges financial support from JSPS KAKENHI Grant-in-Aid for Scientific Research (C) (Grant Number 17K05896). Competing interests The authors declare no competing interests originating from this work. Contributions All authors contributed to the manuscript and its final approval. AK and MF initiated this study, while IVP joined in designing of this study, MF performed the experiments and data analysis. AK and IVP helped with the interpretation of the data and data analysis. AEN, MEC and ZB provided active Gs cells for this study, and participated in discussion together with PB. Dedication The authors dedicate this paper to Prof. Thomas Wandlowski († 2015) for his inspiring guidance. Appendix A. Supplementary data Supplementary data to this article can be found online at https://doi. org/10.1016/j.bioelechem.2018.11.013.

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