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Efficiency of Enzymatic O2 Reduction by Myrothecium verrucaria Bilirubin Oxidase Probed by Surface Plasmon Resonance, PMIRRAS and Electrochemistry Cristina Gutierrez-Sanchez, Alexandre Ciaccafava, Pierre-Yves Blanchard, Karen Monsalve, Marie-Thérese Giudici-Orticoni, Sophie Lecomte, and Elisabeth Lojou ACS Catal., Just Accepted Manuscript • DOI: 10.1021/acscatal.6b01423 • Publication Date (Web): 15 Jul 2016 Downloaded from http://pubs.acs.org on July 24, 2016

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ACS Catalysis

Efficiency of Enzymatic O Reduction by 2

Myrothecium verrucaria Bilirubin Oxidase Probed by Surface Plasmon Resonance, PMIRRAS and Electrochemistry Cristina Gutierrez-Sanchez,[a] Alexandre Ciaccafava,[b] Pierre Yves Blanchard,[a] Karen Monsalve,[a] Marie Thérèse Giudici-Orticoni,[a] Sophie Lecomte, [c] and Elisabeth Lojou*[a] [a]Aix Marseille Univ, CNRS, BIP, UMR 7281, 31 Chemin Aiguier, 13402 Marseille, France. [b]Technische Universität Berlin, Institut für Chemie, Sekretariat PC 14, D-10623 Berlin, Germany [c]Institut for Chemistry and Biology of Membrane and Nanoobjects, Allée Geoffroy St Hilaire, 33600 Pessac, France

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ABSTRACT

Deciphering which parameters control the immobilization of enzymes on solid supports is essential for the development of biotechnological devices such as biosensors, bioreactors or enzymatic fuel cells. In this work, we used Surface Plasmon Resonance (SPR) coupled to electrochemistry and Polarization Modulated Infrared Reflection Absorption Spectroscopy (PMIRRAS) to correlate the loading, the conformation and the activity of Myrothecium verrucaria bilirubin oxidase (Mv BOD) enzymes immobilized on two oppositely charged SelfAssembled-Monolayers (SAMs) on gold electrodes. SPR signal showed that an enzyme layer close to a monolayer was formed by spontaneous adsorption on both negatively and positively charged SAMs. A different catalytic process for O2 reduction was obtained however, being a direct catalysis at negative interfaces and a mediated catalysis at positive interfaces, in relation with the charge of the amino acids surrounding the surface of the Cu T1 and the dipole moment direction of Mv BOD. The stability of the enzymatic current was dependent on the SAM type. On the positively SAM electrode, the mediated catalytic current was stable with time. On the negatively charged SAM, the direct catalytic current decreased continuously with time, leading to a decrease of the TOF (turnover frequency) from 114 to 7 s-1, while the SPR signal remained stable, showing that the decrease in the catalytic current is not related to a desorption process. PMIRRAS studies suggested a conformational change in the tertiary structure as a result of strong electrostatic interactions between arginine residues close to the T1 Cu and the carboxylic functions on the SAM. Covalent binding however resulted in a great enhancement of the current stability, which can be explained by a rigidification of the enzyme layer.


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KEYWORDS. SPR; PMIRRAS; Electrochemistry; Bilirubin oxidase; Direct Electron Transfer; Mediated Electron Transfer.

1. INTRODUCTION The understanding of the main energy metabolic chains implies the knowledge of the affinity between physiologic partners, and the quantification of the reactions rates such as enzymatic reactions, or inhibition reactions. In these chains, when electrons are exchanged between two partners, hydrophobic and electrostatic interactions induce a favorable orientation of one partner against the other to optimize the intermolecular electron transfer rate.1 Electrochemistry is one elegant tool to study these interactions and to obtain some of the main kinetic and thermodynamic parameters involved. Mimicking the physiological environment of the enzyme at the electrochemical interface is expected to provide the suitable conditions for a fast and direct interfacial electron transfer. The success of this concept supposes that the key parameters that allow a favorable orientation of the enzymes are identified. In this respect, the electron relay on the protein surface must be positioned at a distance to the electrode surface allowing fast electron transfer. Thanks to high resolution protein crystallographic structure coupled to accurate modeling and surface spectroscopy, the molecular basis for chemical functionalization of the electrochemical interface for a suitable orientation of some redox enzymes can be drawn.2,3,4,5,6,7Nevertheless, it is admitted that a few percentage of currently purified proteins can be addressed by electrochemistry.8 This clearly illustrates how crucial is the knowledge of the parameters that drive electron transfer processes involving immobilized enzymes.9


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This fundamental achievement has consequences on the development of bioreactors, biosensors, but also biodevices producing electricity. The so-called enzymatic fuel cells (EFCs) nowadays appear as sustainable alternatives to fossil fuels.10,11 In H2/O2 EFCs as an example, the entrapment of hydrogenase and bilirubin oxidase in suitable functionalized 3D matrix induced a drastic increase in the performances, reaching the required power for small electronic devices such as environmental sensors.12,13 However, the applicability of EFCs are limited by their long term stability. In the particular case of H2/O2 EFCs that we are currently developing, a loss between 40 % and 60% of the performance of the EFC during 24 h of continuous operation was reported.12,13,14 Otherwise, in many fundamental electrochemical experiments aiming to elucidate catalytic mechanisms, decrease of the direct catalytic current (DET) reflecting the activity of an enzyme on an electrochemical interface is also observed with time, usually referred as “film loss”. This is however a straightforward idea and there is a requirement to recognize and decipher between the various origins for such instability among protein leaching, reorientation, reconformation, denaturation…, then to be able to propose a remediation process. Unfortunately, non-catalytic signals are rarely observed for enzymes. Moreover, the DET process does not address all the enzymes present at the electrochemical interface, but only those with the surface electronic relay at a short distance from the electrode compatible with the electron transfer. Based on DET electrochemical signals, it is thus hardly possible to link the catalytic activity in various conditions to the amount and structure/conformation of adsorbed proteins. Besides, mediated electron transfer (MET) process is still employed in many biodevices, either because the conditions for DET are unknown or because this is one way to protect the enzyme from external damages.15,16,17,18 One essential question is whether an enzyme electrically connected to an interface would be less stable than an enzyme connected through a


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redox mediator. All these issues require to couple electrochemistry to other surface methods able to quantify and follow the enzyme amount and conformation as a function of the interface environment. The bilirubin oxidase from Myrothecium verrucaria (Mv BOD) is used as a reference model for this study, because the crystallographic structure19 is resolved and many kinetic parameters have been obtained on various electrochemical interfaces.20Mv BOD is a fungal multicopper oxidase that contains four copper atoms. Physiologically, electrons are received from the substrate to the T1 Cu and transferred through a histidine-cysteine bridge to the trinuclear Cu cluster 12-14 Å away where O2 reduction into water takes place. The T1 Cu center situated at 7-8 Å from the surface of the enzyme is targeted for fast interfacial electron transfer rate.21 It was early reported that the modification of graphite electrode by bilirubin or by analogs of bilirubin, the natural substrate of BOD, enhanced the catalytic current for O2 reduction.19,22 This effect was thought to be related to orientation of the enzyme thanks to binding through the substrate pocket. Computational studies further proposed that bilirubin could serve as an electronic extension of the electrochemical interface.23 Some very recent papers efficiently use negatively charged molecules structurally related to bilirubin for carbon nanotube modification.24 Based on the knowledge that a hydrophilic pocket composed of positively charged amino acid residues surrounds the T1 Cu, many other studies also reported that carboxylate groups on the surface of electrodes allow fast electron transfer for adsorbed Mv BOD.21,25,26,27 Although this observation is commonly accepted to be related to a suitable orientation of the enzyme on the surface, the proof of such an orientation, and the way on how conformation and amount of adsorbed proteins evolved in relation with the catalytic activity are only seldom reported.4,28 Coupled methods were used to get further insights in the relation between the enzymatic activity


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and the immobilized enzyme amount and conformation.29,30 Quartz crystal microbalance coupled to electrochemistry (e-QCM) was especially carried out to study Mv BOD activity on negatively charged interfaces.31,

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It was shown that electroactivity loss did not correlate with enzyme

leaching but with structural rearrangements.33,34 Surface Plasmon Resonance (SPR) is another method that can be coupled to electrochemistry in order to obtain information about adsorbed species at the electrochemical interface.35Any perturbation at the surface induces change in the refractive index of the adsorbed layer that is monitored through the modification in the resonant angle of light as a result of light adsorption by surface plasmons.36,37 Electrochemistry coupled to SPR (e-SPR) has been used in the past to study electron transfer on cytochrome c as a function of SAM chemistry on gold electrode,38 redox state-induced immobilization of ferrocene-modified proteins,39 effect of nanostructuration of materials on the activity per bound glucose oxidase molecule,40 assemblies of various nanomaterials and enzymes for biosensor applications.41 In this work, we perform e-SPR to study for the first time the relationship between loading and activity of Mv BOD adsorbed on two different SAMs. The influence of the chemistry of the SAM, the potential at which the catalysis is performed, the presence of coupling reagents on the relationship between catalytic activity and loading and stability of adsorbed proteins is first discussed. Then, the MET process on the two different SAMs is especially analyzed in order to address the question of the stability and conformation of the enzymes oriented for a DET or a MET process. Polarization Modulated Infrared Reflection Absorption Spectroscopy (PMIRRAS) is carried out to give access to the conformation (secondary structure) and the orientation of the protein adsorbed on the different modified electrodes. Spectroscopic data are further discussed with complementary analysis of Mv BOD structure, allowing us to propose a model for Mv BOD orientation on the electrode depending on the type of SAMs. To


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the best of our knowledge, this is the first time adsorption of Mv BOD is studied by e-SPR with a correlation of the evolution of the catalytic signal, both DET and MET. The results obtained here provide the experimental tools to rationalize electrochemical interfaces suitable for fast electron transfer processes.

2. METHODS 2.1. Chemicals and materials Mv BOD was a gift from Amano Enzyme Inc. (Nagoya, Japan). Fresh solution of Mv BOD was prepared in 100 mM phosphate buffer at pH 6. The concentration of Mv BOD was measured at 595 nm by spectrophotometry using Bradford test with BSA standard (the molecular weight of Mv BOD is 60 kDa). The purity of the enzyme was checked by SDS gel. Ethanol analytical grade 96% (v/v), 4-aminothiophenol (ATP), 6-mercaptohexanoic acid (MHA), 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide (EDC), N-hydroxysuccinimide (NHS), 2,2'-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS), hydrogen peroxide solution 30 % (H2O2), sodium fluoride 99 % (NaF), sodium hydroxide 97 % (NaOH) and sulfuric acid 95-98 % (H2SO4) were purchased from Sigma-Aldrich. 2.2. Electrochemistry measurements Electrochemical experiments were performed at room temperature using a potentiostat from Autolab PGSTAT30 analyzer controlled by GPES 4.9 software and Nova software (Eco Chemie). The electrochemical cell, which corresponds to the SPR cuvette, with a volume of 300 µL was equipped with three electrodes, the SPR gold disc as a working electrode with a surface of 0.07 cm2, a platinum wire as an auxiliary electrode and an Ag/AgCl electrode as a reference


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electrode. Standard deviation for the catalytic current was calculated from three measurements using different electrodes. 2.3. SPR measurements SPR measurements were acquired on Autolab SPRINGLE instrument (Eco Chemie, The Netherlands) by using bare planar gold discs (25 mm diameter) purchased from Eco Chemie. A polarized laser light (λ=670 nm) was directed to the bottom side of the sensor via a hemispheric lens placed on a prism (BK7 with a refractive index of 1.52) and the reflected light was detected using a photodiode. The gold disk was mounted on the hemispheric lens to form the base of the cuvette. For the SPR measurements, 100 µl of 0.1 M phosphate buffer, pH 6 was injected into the cell until stabilization of the signal was achieved. The buffer solution was then replaced by a solution of 25 µM Mv BOD (1.5 mg/mL) in buffer and the SPR signal was monitored to follow enzyme adsorption. At the end of the adsorption process, enzymes remaining in solution and loosely adsorbed molecules were removed out from the cell by buffer washing. The procedures for sample injection and removal were carried out using an autosampler (EcoChemie) equipped with a peristaltic pump. Data were analyzed by a SPR software from EcoChemie. The mass of the adsorbed species was calculated from the SPR signal based on the relation that a change of 122 mdeg (millidegrees) corresponds to 1.0 ng/mm2 at 25 °C.42,43,44 At least five experiments were conducted, and only the electrodes whose surface coverage deviation did not exceed 10 % from the average were used. Standard deviation was calculated from the measurements using different electrodes.


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2.4. Electrode preparation The gold substrates were cleaned with “piranha” solution (3:1 H2SO4:H2O2) (CAUTION: Piranha solution is especially dangerous, is corrosive, and may explode if contained in a closed vessel; it should be handled with special care) during 4 min and rinsed extensively with water and later with ethanol. SAMs were formed by immersing the gold electrodes in 5 mM ethanol solutions of ATP or MHA during 18 hours. The surface was then cleaned with ethanol to remove all organic contaminants. These SAM gold electrodes, named ATP-Au and MHA-Au, respectively, were placed onto the prism of SPR system to monitor simultaneously the SPR signal and the electrochemical signal. In the case of covalent binding, 50 µL of 36 mM EDC and 50 µL of 17 mM NHS solution were added and let react during 15 min with the enzyme layer, before rinsing and running the electrochemical experiment. 2.5. PMIRRAS measurements For PMIRRAS, the modified dried gold electrode was placed at room temperature in the external beam of FT-IR instrument on a Nicolet Nexus 870 FT-IR spectrometer (Madison, WI), and the reflected light was focused on a nitrogen cooled (77K) HgCdTe (MCT) detector (SAT, Poitiers). The optimal value of the angle of incidence for the detection was 75° relative to the optical axis normal to the interface. A ZnSe grid polarized and a ZnSe photoacoustic modulator to modulate the incident beam between p and s polarizations are placed prior the sample. The detector output is sent to a two-channel electronic device that generates the sum and the difference interferograms. The PMIRRAS spectra were recorded at 8 cm-1 resolution, with coaddition of six hundred scans. Using a modulation of polarization enabled us to perform rapid analyses of the sample after treatment in various solutions without purging the atmosphere or requiring a reference spectrum.


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2.6. Structure Analysis Mv BOD structure (pdb entry: 2XLL) was analyzed and images were created with Pymol (The PyMOL Molecular Graphics System, Version 1.7 Schrödinger, LLC). The dipole moment of Mv BOD was computed online using the Protein Dipole Moments Server available at (http://dipole.weizmann.ac.il/).

3. RESULTS AND DISCUSSION 3.1. Adsorption of Mv BOD on the negatively charged MHA-Au surface Enzyme adsorption on MHA-Au surface was first quantified using SPR. Mv BOD was adsorbed at open circuit voltage (OCV) during 2 h, then the surface was rinsed with buffer. A typical SPR signal is given in Figure 1. Before Mv BOD injection at t = 0, the SPR signal recorded in buffer is stable and is taken as the baseline. The SPR signal increases during the Mv BOD adsorption phase and reaches a plateau after ca. 4000 s indicating the adsorption equilibrium. The rinsing step induces a small decrease of the signal due to some Mv BOD molecules weakly adsorbed. This amount is evaluated to be less than 5 % of the total adsorbed amount. A difference of 350 mdeg is observed between the value obtained just before injection of 25 µM Mv BOD and after rinsing. The angle variation corresponds to a surface coverage of adsorbed Mv BOD, ΓSPR of 4.8 ± 0.7 pmol.cm-2 which has to be compared to the theoretical surface coverage Γth of a densely packed Mv BOD. Depending on the orientation of the enzyme based on the geometrical dimension of Mv BOD (40x50x60 Å) obtained from the X-ray crystallographic structure,19,45 Γth is between 5.5 to 8.3 pmol.cm-2. The experimental Mv BOD surface coverage is thus in the order of a monolayer in agreement with previous values obtained with Mv BOD adsorbed on SAMs containing carboxylate as a head group.29


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The kinetics of adsorption has been fitted with a biphasic model interaction (details of the fit is given in SI, and the kinetic parameters are summarized in table S1), in agreement with the suggestion that the electrode area equivalent to the footprint of the protein molecule should include more than one amino acid residue to achieve an efficient adsorption of the protein.46

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Figure 1. SPR measurement for Mv BOD adsorption on MHA-Au at OCV in 100 mM airsaturated phosphate buffer, pH 6.0. 25 µM Mv BOD was injected at t = 0 s. Fit of the kinetic SPR curve is represented in blue dots. Inset: CVs recorded at t1 (black line) and t2 (red line) during Mv BOD adsorption on MHA-Au at OCV in air-saturated 100 mM phosphate buffer pH 6.0. The grey line corresponds to the CV signal obtained after 30 mM NaF injection at the end of the adsorption process. v = 0.01 V.s-1.

3.2. Direct electrocatalytic reduction of O2 by Mv BOD on the negatively charged MHA-Au The aforementioned electrode was characterized by cyclic voltammetry performed directly in the SPR cuvette. Inset in Figure 1 gathers the typical cyclic voltamograms (CVs) recorded after 3


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min (black line) and 90 min (red line) of Mv BOD adsorption, corresponding to times t1 and t2 on the SPR signal (Figure 1), respectively. For both adsorption times, a reductive catalytic wave starts at a potential close to the redox potential of Mv BOD Cu T1 site.47 NaF completely vanishes the catalytic signal (grey dotted line) as expected for inhibition of the catalytic activity because fluoride binding to the T2/T3 Cu cluster.48,49 At t2, an increase of 30 % of the signal was observed when O2 was bubbled inside the buffer (data not shown).

The enzymatic origin of the reduction process is thus demonstrated. It involves Mv BOD molecules oriented at the electrochemical interface with the Cu T1 at a distance compatible with DET. MHA has a pKa of 4.3.50 The Mv BOD structure displays a dipole moment of 752 Debye pointing toward the highly positive patch formed by the 4 arginine (R353, R356, R436 and R437) (pKa = 12.5) residues surrounding the T1 Cu center (Scheme S1). Although the Debye length (close to 1 nm) is shorter than the radius of Mv BOD, such an environment clearly promotes DET on MHA-Au which is negatively charged at pH 6. A much higher DET current density is recorded at t1 (14 µA.cm-2) than at t2 (8 µA.cm-2). However, the corresponding surface coverage at t1 and t2 calculated from the SPR signal are 0.6 and 5 pmol.cm-2, respectively. From the limiting catalytic current and the amount of adsorbed Mv BOD on MHA-Au electrode, a turn over frequency kcat of 114 s-1 and 7 s-1 are calculated (see details of the calculation in SI), respectively at t1 and t2. kcat at short adsorption time is in accordance with previous value obtained for pure O2 reduction on a rotating disc electrode,51 while kcat at longer adsorption time is in the same order of magnitude than previous reported values obtained from Mv BOD immobilized on gold substrates.30,33,34Our results underline that the bioelectrode modified during short times presents a much better enzymatic activity per


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number of grafted enzymes. We further confirmed this observation by running a second set of experiments with Mv BOD at a lower concentration of 1 µM (Figure S1). The surface coverage was far less than a monolayer, typically 0.6 pmol.cm-2. But the catalytic current takes a value of 6 µA.cm-2, yielding kcat of 48 s-1. The decrease in the activity relative to the amount of adsorbed proteins suggests that a great proportion of adsorbed enzymes is not electroactive upon immobilization, but also that the proteins already adsorbed tend to be inactivated with time. This conclusion can be connected to results by Taniguchi and coworkers who compared the surface coverage obtained from QCM and the surface coverage of electroactive Mv BOD obtained from CV.29 They showed that only ca. 10% of the adsorbed enzymes were electroactive. Our results would also suggest that a submonolayer would be more catalytically efficient than the monolayer. In relation with this hypothesis, Blanford et al. 51reported that there was an optimized concentration of BOD to enhance the catalytic activity and stability. Noteworthy, the catalytic currents in this latter work were ten times higher than in the present study and obtained for a ten times higher BOD concentration. However, BOD concentrations used in the present work and the value of current recorded are in the same range than observed in other published works with Mv BOD immobilized on gold electrodes under O2.30,33,34 Control experiments in our experimental conditions did not show an increase neither in the surface coverage nor in the direct catalytic current by using BOD concentration of much higher concentration (i.e. 250 µM, data not shown). These discrepancies may originate from different experimental conditions, i.e. type of the SAM and especially the length of the thiol, catalysis under air against O2-saturated buffer, geometric surface of the electrode vs volume of the SPR cell. To further analyze the activity loss, we chose an adsorption time of Mv BOD of 15 min before rinsing with buffer, which appeared as


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a good compromise between maintaining reproducible activity and approaching the plateau for Mv BOD adsorption. The DET evolution with time was then studied by running a CV experiment every 1000 s during 4000 s after the steps of Mv BOD adsorption then rinsing, and the SPR signal was simultaneously recorded (Figure 2). After 15 min of Mv BOD adsorption then rinsing, the SPR angle allows evaluating a surface coverage of 4.1 ± 0.6 pmol.cm-2. A very stable SPR signal is observed, showing a variation of only 1.5 mdeg over the time of experiment, indicating a high stability of the loading of Mv BOD. The DET current for the reduction of O2 continuously decreased with successive CVs, however. The total current decrease is evaluated to be around 40 %. kcat consequently decreases from 13 s-1 to 7 s-1. The calculated kcat are obtained from the surface coverage. Their values reflect first that all enzymes are not catalytically active as already noticed from the comparative catalytic currents obtained during Mv BOD adsorption phase. Their evolution as a function of time reflects a loss of activity of the grafted enzymes. Addition of 25 µM fresh Mv BOD in the SPR cuvette after 4000 s did not allow recovering of the initial catalytic current, confirming that the decrease in the catalytic current cannot origin from a simple loss of protein from the electrochemical interface.


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Figure 2. SPR signal recorded for 15 min during 25 µM Mv BOD adsorption on MHA-Au, then during 4000 s after rinsing. Inset: CVs recorded every 1000 s at the times CVi pointed on the SPR curve in air-saturated 100 mM phosphate buffer pH 6.0. v = 0.01 V.s-1. The grey line corresponds to the CV signal obtained after 30 mM NaF injection at the end of CV5. The coupling between SPR and electrochemistry clearly demonstrates that the decrease in the catalytic current is not related to protein loss but mainly from a non-desorption phenomenon. We have checked that the decrease in the catalytic signal is not attributable to the spontaneous deactivation of the enzyme. Indeed, a CV was recorded using Mv BOD incubated during 4000 s in a buffer solution at room temperature before being adsorbed on MHA-Au. A catalytic signal similar to that one obtained at CV1 of Figure 2 inset was obtained (Figure S2). This means that only the immobilized state induces a decrease in the enzyme activity during the time of the experiment. The decrease in the catalytic signal is neither coming from O2 depletion. Bubbling O2 directly in the cell after CV5 of Figure 2 inset did not allow to recover the initial catalytic signal (not shown).


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One question is whether the instability of the catalytic current could be related to the specific orientation of enzyme required for a direct electron transfer process. The electrostatic interactions involved in the recognition process could induce a change in the conformation of the protein. In this case, it should be expected that applying a potential would also impact the catalytic signal stability.31,32The same experiment as described in Figure 2 was repeated but poising the Mv BOD modified MHA-Au at a potential of +0.2 V vs Ag/AgCl during 4000 s. CV experiments were again recorded every 1000 s while the SPR signal was simultaneously followed (Figure 3).

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CV Figure 3. A) SPR measurement recorded for 15 min of 25 µM Mv BOD adsorption on MHAAu. At time denoted CAi on the SPR signal, a potential of +0.2 V vs Ag/AgCl was applied. At times denoted CVi, a CV was recorded and is reported in inset. The grey line corresponds to the CV signal obtained after 30 mM NaF injection at the end of CV5. Air-saturated 100 mM phosphate buffer pH 6.0. v = 0.01 V.s-1. B) Comparison of the catalytic current loss at OCV (red columns) or under +0.2 V applied potential (black columns). The error bars are represented in green.

The SPR signal is more affected than when recorded at OCV. A slight decrease in the SPR angle after 4000 s is observed which corresponds to 3 % of the initial signal. On the other hand, a high decrease in the catalytic current corresponding to 70% of the initial current is measured. The coupling between SPR and electrochemistry under an applied potential again leads to the main conclusion that the catalytic current decrease is not related to protein release from the


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surface. The total percentage of activity loss is otherwise strongly dependent on applying a potential or not. However, as underlined in the histogram in Figure 3B, the loss in catalytic current mainly occurs between CV1 and CV2. It is twice higher under applied potential than at OCV. The percentage of catalytic loss is afterwards more similar under applied potentials or at OCV, although still more marked under applied potential. This suggests that applying a potential may induce a first step of enzyme layer organization which leads to enzyme deactivation.

3.3. Mediated O2 reduction by Mv BOD on the negatively charged MHA-Au It can be hypothesized that the decrease in the direct electrocatalytic current may be related to reorientation of the protein with time and/or under potential. To address this issue, ABTS, a typical redox mediator for Mv BOD, is added in solution to evaluate the mediated electrical connection of potentially unfavorably reoriented enzymes. When ABTS is added in condition similar to CV1, it induces a very small variation of the DET signal, showing that all the enzymes are orientated to produce DET (Figure 4A). When ABTS is added after DET current has almost vanished, no recovery of the catalytic signal can be observed (Figure 4B). From the subtraction between the remaining DET current and the current in presence of ABTS, only the signal relative to the redox behavior of ABTS alone can be obtained (Inset in Figure 4B). This result demonstrates that the activity loss is not due to a modification in the orientation of Mv BOD at the interface. It could be otherwise argued that the enzymes not oriented in DET would be more stable, because less impacted by the electric field. This is an important open question in the development of stable biodevices such as biofuel cells. It was shown as examples that the integration of soluble hydrogenases in viologen modified redox hydrogels or in carbon nanotube-


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polypyrrole films allows the enzymes to be protected from deactivation at high potentials.52,16 This protective role of the mediated connection was not so clear for membrane-bound [NiFe]hydrogenases which oxidize H2 at higher potentials, then use higher redox potential mediators.16 To study the stability of the MET process with time, the adsorption time for Mv BOD was increased to 90 min (Figure 4C). CVs were first performed before and after ABTS addition just after the rinsing step. The resulting subtracted CV (green line in Figure 4C) presents a sigmoidal shape clearly indicating the occurrence of some MET process. The proportion of MET current can be evaluated to 21%, and can be explained by the increase in the surface coverage which induces an increase in the distribution of orientation. The bioelectrode was then let at OCV during 2000 s, and the CVs before and after ABTS addition were performed again. The resulting subtracted CV (dashed green line in Figure 4C) corresponds to the signal for ABTS alone. These last experiments demonstrate that even the enzymes not in direct connection with MHA-Au do not provide a stable electrocatalytic signal.

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Figure 4. CV studies of the MET process on MHA-Au. (A) CV as CV1 in Figure 3 (black line) and after 10 µM ABTS addition (green line); B) CV as CV5 in Figure 4 (black line) and after 10 µM ABTS addition (green line); inset of (B) corresponds to the CV subtraction; (C) MET contribution after 90 min Mv BOD adsorption just after rinsing (green line) and after 2000 s (dashed green line) at OCV. v = 0.01 V.s-1


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3.4. Mediated O2 reduction by Mv BOD on the positively charged ATP-Au The experimental data obtained in this work highlight deleterious effect induced by carboxylate groups present on the electrochemical interface on Mv BOD activity. One can expect different stability responses according to the functionality on the electrode. Kamitaka et al. concluded that a hydrophilic surface was required to avoid protein denaturation.34 Sugimoto et al. demonstrated that a hydrophobic SAM layer prevents the electric field to affect the protein adsorption process.31 e-SPR was thus used in this work to study the stability of Mv BOD adsorbed on interfaces bearing other chemical groups than COO-. The SPR gold electrode was modified with ATP, and Mv BOD was adsorbed on the ATP-Au electrode. Actually, the pKa of 4-ATP is 6.9,53 thus yielding a positively charged electrochemical interface at pH 6. Mv BOD is thus expected to be preferentially oriented for a MET process. The corresponding SPR and electrochemical responses are given in Figure 5. Mv BOD is adsorbed on ATP-Au following a biphasic kinetics (Table S1). A 335 mdeg difference is observed before and after Mv BOD adsorption at t2 (8000 s). This angle corresponds to a surface coverage of 4.5 ± 0.6 pmol.cm-2, very close to a monolayer. As can be seen in the inset of Figure 5, the electrochemical response of the modified bioelectrode does not present any DET process, in contrast to the response obtained on MHA-Au. Nevertheless, reduction of O2 occurs after ABTS addition in the SPR cell, thus showing that molecules of Mv BOD attached to the electrode interface were unable to take electrons directly from the gold electrode because of unfavorable orientation. The value of the limiting current in this case (close to 10 µA.cm-2) is used to calculate the pseudo-first order rate constant kf to 15 s-1.54


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Figure 5. SPR measurement recorded for 25 µM Mv BOD adsorption on ATP-Au during 8000 s. Fit of the kinetic SPR curves is represented in blue dots. Inset: CVs of O2 reduction by Mv BOD adsorbed on ATP-Au electrode at t2 before (black lines) and after 10 µM ABTS addition (green line). The gray line corresponds to the CV in the presence of 30 mM NaF after ABTS addition. 100 mM air-saturated phosphate buffer pH 6.0, v = 0.01 V.s-1. To compare the stability of MET process on ATP-Au compared to DET and MET on MHAAu, CV experiments were performed on Mv BOD-modified ATP-Au at times t’1 and t’2 on the SPR curve obtained after 15 min Mv BOD adsorption. The corresponding SPR signal and CVs are presented in Figure 6.


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Figure 6. A) SPR measurement recorded for 25 µM Mv BOD adsorption on ATP-Au for 15 min, then after rinsing for 4000 s; B) CVs obtained at times t’1 (red curve) and t’2 (blue curve) on the SPR curve in the presence of 10 µM ABTS. The CVs are obtained after subtraction of the capacitive current. 100 mM air-saturated phosphate buffer pH 6.0, v = 0.01 V.s-1


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The SPR signal is stable with time, displaying a variation less than 10 mdeg, i.e. 4% of the total signal after 4000 s. As shown on Figure 6B the MET process is only slightly affected by the time of storage of BOD-ATP-Au. Mediated catalytic current densities respectively of 4.9 and 4.6 µA.cm-2 are obtained respectively at t’1 and t’2. The MET process is thus much more stable when Mv BOD is adsorbed on NH2-terminated surface than on COOH-terminated surface, suggesting different interaction.

3.5. Conformation and orientation of Mv BOD adsorbed on negatively charged MHA-Au and positively ATP-Au SPR and electrochemistry experiments alone are not sufficient to identify the forms of the inactivated or denatured enzymes. To assess any conformational or orientational change of Mv BOD depending on the chemistry of the surface, PMIRRAS has been performed on MHA-Au and ATP-Au modified by the enzyme (Figure 7). Figures 7A and 7B show the spectra recorded after Mv BOD adsorption on the MHA-Au or ATP-Au, respectively. Mv BOD adsorption on the modified gold electrode is confirmed by the presence of the amide I (mainly C=O stretching vibrational mode) and the amide II (mainly N-H stretching vibrational mode) at around 1663 and 1540 cm-1, respectively. Some additional bands (marked with stars on Figures 7A and 7B) are observed. They are characteristic of the SAM used for adsorption. The bands at 1729 cm-1 and 1589 cm-1 are assigned to carbonyl group vibrations of the MHA SAM and to C=C aromatic stretching mode of ATP, respectively (Figures 7A, 7B and Figure S3). Similar PMIRRAS spectra are observed just after the immobilization or after 1 hour after the adsorption (results not shown), suggesting no modification of the secondary structure with time.


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For all modified electrodes, PMIRRAS spectra present the amide I at the same wavenumber, however at a higher position compared to the ATR-FTIR spectrum of Mv BOD in solution (Figure S3). This increase of around 20 cm-1 could be due to secondary structure modification or to specific orientation of the enzyme adsorbed on the modified electrodes. From the Mv BOD ATR-FTIR spectrum, we evaluated the proportions of amide groups involved in the secondary structure elements. 34% of the amide groups are involved in β-sheet, 19 % in α-helices and 23 % in random coil configuration, in agreement with the elements of secondary structure observed by crystallography (details are given in supplementary materials Table S2). The appearance of the bands at 1661 cm-1 on the PMIRRAS spectra could reveal a large modification of the secondary structure of Mv BOD implicating a large increase of the amide groups involved in α-helices at the expense of the β-sheets and random coil. However, since the Mv BOD is still catalytically active after adsorption on modified electrodes, strong secondary structure modification is rather improbable. Olejnik et al. reported a similar shift for FTIR spectra of solid laccase recorded in the KBr pellet compared to the PMIRRAS spectra of laccase immobilized on thiol modified gold electrode, and draw similar conclusion. The authors also observed a supplementary band at 1733 cm-1 that they attributed to protonation of key glutamic residues involved in O2 reduction. In our experiments, however, the band at 1729 cm-1 appears only on MHA-Au, and arises from the SAM layer itself (Figure S4). Due to the surface selection rule for infrared reflectance, only vibrations having component perpendicular to the gold surface contribute to the spectrum.4,55,56 Then the low intensity observed on the PMIRRAS spectra of β-sheets and random contributions at 1630-1643 cm-1 can be only due to the specific orientation of β-sheets tilted on the surface. The orientation of αhelices seems to be more perpendicular to the surface allowing a higher contribution in the


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PMIRRAS spectra. Unexpectedly the amide I/amide II band area ratios are similar whatever the surface MHA or ATP used for the immobilization of the enzyme. It seems to indicate that both the secondary structure and the orientation of the enzyme are identical for adsorption on MHA or on ATP surface. However, the catalytic activities are radically different (see above), as the electron transfer processes either by DET or by MET. This apparent discrepancy can be rationalized by a closer visual inspection of the Mv BOD crystallographic structure, taking into account the topology and the type of secondary structures in addition to the calculated dipole moment of the enzyme. As laccases, Mv BOD topology consists of a three-fold repeated β-barrel motif. Those three β–barrels run nearly parallel to each other, and the dipole moment of the protein lies perpendicular to them. The implication is straightforward; most of the β-sheets shaping the barrels are perpendicular to the dipole moment. Consequently, depending on the charge of the SAM used to modify the electrode surface, flipping the enzyme upside down along the dipole moment results roughly in the same projected orientation and amount of β-sheets on the surface (Figures 7C and 7D). Moreover, the orientation of α-helices is conserved after this rotation. This hypothesis is also consistent with the low amount of observed β-sheets. Hence, it is not unexpected to observe similar spectral feature whereas the electrochemical behavior is drastically different. To explain the decrease with time of the catalytic activity on MHA-Au, we may suggest that it is more related to modification of the enzyme tertiary structure or to modification surrounding the T1 Cu center. Strong interactions can effectively occur between the lateral chain of the arginine residues and the COO-group of the MHA monolayer. These interactions will slightly modify the tertiary structure of the enzyme close to the redox center explaining the decrease of


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the catalytic activity. The peculiar stability that we observe for MET process on ATP-Au may then origin from softer interaction with the enzyme.

Figure 7. A) PMIRRAS spectrum of Mv BOD adsorbed during 15 min on negative MHA-Au. B) PMIRRAS spectrum of Mv BOD adsorbed during 15 min on positive ATP-Au. Corresponding schematic representation of the Mv BOD orientation along its dipole moment depending on the charge of the SAM modified electrode on MHA-Au C) and on ATP-Au D). Mv BOD secondary structures are depicted in different colors, β-sheets in red, α-helices in green and random coil in purple. Random coil has been removed from the 90° rotated profile view in order to increase visibility of the β-sheets. The dipole moment is represented by an arrow. Mv BOD structure from pdb entry: 2XLL.


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3.6. Increasing of the stability of negatively charged MHA-Au biolectrode by covalent coupling Covalent bonding can be used to enhance the stability of bioelectrodes, including multicopper oxidase-based ones. This was previously done by cross coupling of BOD and the electrode using glutaraldehyde,57,58 or maleimide derivatives,59 or EDC/NHS coupling.26,60 Enhancement of stability by covalent attachment was reported on other enzymes such as cellobiose dehydrogenase or hydrogenases.58,61 De Lacey et al. especially showed that the operational stability of hydrogenase directly adsorbed on the electrode was very low, whereas the electrodes with hydrogenase immobilized by covalent bond were very stable after a month of continuous operation.62 Conversely, it was reported that covalent attachment induced a slower electron transfer rate for O2 reduction by Mv BOD, and can even prevent any catalytic signal.30,32 The discrepancy between experimental data may origin from the type of enzyme, but also from the way the covalent attachment is realized as we will detail in the following part. In this work, we demonstrate the absence of release of protein with time, and so we expected that EDC/NHS would have no impact on the stabilization of the catalytic signal with time. To check this assumption, EDC/NHS was used to generate an activated ester and then form an amide bond between the COO- of the MHA-Au and the NH2-amino acid residues (arginine residues (R353, R356, R436 and R437) located approximately 13 Å from T1 center, Scheme S1). To achieve this experiment, Mv BOD was first oriented on the MHA-Au to get a proper orientation of the enzyme, then EDC/NHS (2:1) mixture was added to react with the Mv BODmodified MHA-Au. Adsorption, then stability, of Mv BOD was followed by SPR and electrochemistry measurements in the same cell (Figure 8).


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E / V vs. Ag/AgCl Figure 8. (A) SPR measurement recorded for 25 µM Mv BOD adsorption on MHA-Au in presence of EDC (36 mM) and NHS (17 mM). EDC-NHS coupling agent was added at the time denoted EDC-NHS on the SPR signal, let to react during 15 min, afterwards the cell was rinsed with buffer; (B) Chronoamperometry at +0.2 V vs Ag/AgCl performed after the coupling step (blue line) (for comparison the chronoamperometry in the absence of the coupling step is given (black line)); (C) CVs obtained at times denoted CV1 and CV2 on the SPR curve before (black line) and after chronoamperometry (red line). 100 mM air-saturated phosphate buffer pH 6.0, v = 0.01 V.s-1

The coupling reaction does not induce any variation with time in the loading of Mv BOD as shown by the value of the SPR signal after rinsing (Figure 8A). Figure 8B compares the evolution of the catalytic current at +0.2 V vs Ag/AgCl with and without the coupling step. A very different stability is observed. A remaining current of only 20% after 4000 s in the absence of EDC-NHS is observed. Conversely, covalent coupling allows 82 % of the initial current to be


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recovered. Moreover, most of the current loss is observed after the first 500 s of chronoamperometry. Although the initial decrease in the current includes a transition step to steady-state, Mv BOD molecules that did not form covalent bonds and were only chemisorbed on the surface, thus more affected by the application of a potential may be also involved. After this initial decay the catalytic current is quite stable. CVs have been performed before and after 4000 s at +0.2V vs Ag/AgCl (Figure 8C) applied to the bioelectrode that has undergone a coupling step. In both cases no MET is observed. At the end of the chronoamperometric step, the CV DET current is still 87 % of the initial CV current, which is incomparably higher than the value recorded in the absence of any covalent bonding (i.e. 30% in Figure 3). From this experiment, two main conclusions can be drawn. First of all, the addition of a coupling agent is not deleterious for the catalytic activity of the enzyme. Secondly, the electrode that has undergone a post treatment with EDC/NHS presents a better catalytic stability than without treatment. Because DET decrease is not due to leaching, we propose that the formation of a covalent link between the pre-oriented BOD and the electrode allows to preserve the suitable orientation for DET and to avoid enzyme denaturation. This behavior is not in agreement with previous results on Mv BOD immobilized on carboxylic-SAM electrodes, which presented a loss of catalytic activity after covalent coupling. The difference can come from the different experimental conditions used in the two studies. Blanford et al.32 immobilized first the activated ester to the gold surface then added the enzyme. Covalent coupling could occur only between the ester of the SAM and NH2 residues from the enzyme with no preferred orientation. In our case, the BOD is already on the surface with the optimized orientation and it may be possible to have some cross-coupling reaction between enzyme and the modified electrode which stabilizes Mv


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BOD DET activity, as proved for other proteins. 62,63 It can be suggested that a suitable enzyme immobilization through multipoint covalent attachments produce a strong rigidification of the enzyme structure favorable to an enhanced stability of the activity.64 We might consider here that covalent attachment should prevent Mv BOD conformational change because of a decrease in the attractive electrostatic interaction following the coupling reaction. We cannot however exclude that covalent bonds take place between amine and acid groups on neighboring proteins resulting also in increasing stability.

4. CONCLUSIONS

In this work, adsorption of bilirubin oxidase from Myrothecium verrucaria on both negatively and

positively charged

electrochemical

interfaces

was

followed

coupling

SPR

to

electrochemistry. SPR showed that the same loading of enzyme is obtained whatever the charge of the electrode, but electrocatalysis proceeds via a DET process on COO- surface against MET process on NH3+, directly in relation with the dipole moment of the enzyme and the amino acid environment of the T1 Cu. The SPR signal was very stable with time while the catalysis continuously decreased, showing that the widely used term “film loss” cannot account for desorption processes. PMIRRAS showed that there is no change in the secondary structure upon enzyme immobilization. The different orientations demonstrated by electrochemistry could not be readily associated with two different orientations by PMIRRAS, although analysis of the crystallographic structure provided consistent reasons to evaluate that in our case, due to a peculiar structural pattern in Mv BOD involving three β-barrels and PMIRRAS selection rules, identical amide I/II ratio does not necessarily mean identical orientation. Nevertheless, covalent


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bond allows stabilization of the catalytic process most probably because of a rigidification of the enzyme layer preventing change in the tertiary structure. We also demonstrated by coupling SPR with electrochemistry that sublayers of enzymes are more active than a compact monolayer. This result suggests that mixed SAMs should be more efficient than pure SAMs. Work is in progress in the laboratory to explore this way for efficient biocatalysis.

AUTHOR INFORMATION Corresponding Author *E-mail for Elisabeth Lojou : [email protected] Author Contributions C.G.S has done the SPR and electrochemistry experiments. S.L. has performed the spectroscopy measurements (ATR-FTIR and PMIRRAS). A.C. evaluated the Mv BOD structure and constructed the orientation model. E.L is the initiator and director of the project and participated in all steps. K.M., P.-Y B. and M-T. G.-O. participated to the discussion of the results. The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. Funding Sources The authors thank ANR for financial support (CAROUCELL-ANR-13-BIOME-0003-02). This work was also supported by A*MIDEX Marseille (ANR-11-IDEX-0001-02). Notes The authors declare no competing financial interest


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ASSOCIATED CONTENT Supporting Information. Additional information about details of the adsorption fit (equation and parameters), equation to calculate kinetic constant, electrochemical measurements, dipole moment of the enzyme, ATR-FTIR spectrum and structural assignments, PMIRRAS spectra of SAM layers. This material is available free of charge via the Internet at http://pubs.acs.org.

ACKNOWLEDGMENT Dr Hafsa Korri-Youssoufi, Paris-Saclay University is especially thanked for helpful discussions on the SPR equipment. The authors are grateful to Marianne Guiral and Marianne Ilbert (BIP, Marseille, France), for fruitful discussion

REFERENCES

(1)

Roger, M.; de Poulpiquet, A.; Ciaccafava, A.; Ilbert, M.; Guiral, M.; Giudici-Orticoni, M. T.; Lojou, E. Anal. Bioanal. Chem. 2014, 406, 1011–1027.

(2)

Rüdiger, O.; Abad, J. M.; Hatchikian, E. C.; Fernandez, V. M.; De Lacey, A. L. J. Am. Chem. Soc. 2005, 127, 16008–16009.

(3)

Ciaccafava, A.; Infossi, P.; Ilbert, M.; Guiral, M.; Lecomte, S.; Giudici-Orticoni, M. T.; Lojou, E. Angew. Chem. Int. Ed. Engl. 2012, 51, 953–956.

(4)

Olejnik, P.; Palys, B.; Kowalczyk, A.; Nowicka, A. M. J. Phys. Chem. C 2012, 116, 25911–25918.

(5)

Oteri, F.; Ciaccafava, A.; de Poulpiquet, A.; Baaden, M.; Lojou, E.; Sacquin-Mora, S. Phys. Chem. Chem. Phys. 2014, 16, 11318–11322.

(6)

Heidary, N.; Utesch, T.; Zerball, M.; Horch, M.; Millo, D.; Fritsch, J.; Lenz, O.; Von Klitzing, R.; Hildebrandt, P.; Fischer, A.; Mroginski, M. A.; Zebger, I. PLoS One 2015, 10, e0143101.


34

Page 35 of 38

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(7)

Ulyanova, Y.; Babanova, S.; Pinchon, E.; Matanovic, I.; Singhal, S.; Atanassov, P. Phys. Chem. Chem. Phys. 2014, 16, 13367–13375.

(8)

Salaj-Kosla, U.; Scanlon, M. D.; Baumeister, T.; Zahma, K.; Ludwig, R.; Ó Conghaile, P.; MacAodha, D.; Leech, D.; Magner, E. Anal. Bioanal. Chem. 2013, 405, 3823–3830.

(9)

Reid, R. C.; Jones, S. R.; Hickey, D. P.; Minteer, S. D.; Gale, B. K. Electrochim. Acta 2016, 203, 30–40.

(10)

Rasmussen, M.; Abdellaoui, S.; Minteer, S. D. Biosens. Bioelectron. 2016, 76, 91–102.

(11)

de Poulpiquet, A.; Ranava, D.; Monsalve, K.; Giudici-Orticoni, M.-T.; Lojou, E. ChemElectroChem 2014, 1, 1724–1750.

(12)

Xu, L.; Armstrong, F. A. RSC Adv. 2015, 5, 3649–3656.

(13)

Monsalve, K.; Mazurenko, I.; Lalaoui, N.; Le Goff, A.; Holzinger, M.; Infossi, P.; Nitsche, S.; Lojou, J. Y.; Giudici-Orticoni, M. T.; Cosnier, S.; Lojou, E. Electrochem. commun. 2015, 60, 216–220.

(14)

De Poulpiquet, A.; Ciaccafava, A.; Gadiou, R.; Gounel, S.; Giudici-Orticoni, M. T.; Mano, N.; Lojou, E. Electrochem. commun. 2014, 42, 72–74.

(15)

Suzuki, A.; Murata, K.; Mano, N.; Tsujimura, S. Bull. Chem. Soc. Jpn. 2016, 89, 24–26.

(16)

Plumeré, N.; Rüdiger, O.; Oughli, A. A.; Williams, R.; Vivekananthan, J.; Pöller, S.; Schuhmann, W.; Lubitz, W. Nat. Chem. 2014, 6, 822–827.

(17)

Ó Conghaile, P.; Falk, M.; MacAodha, D.; Yakovleva, M. E.; Gonaus, C.; Peterbauer, C. K.; Gorton, L.; Shleev, S.; Leech, D. Anal. Chem. 2016, 88, 2156–2163.

(18)

Leech, D.; Kavanagh, P.; Schuhmann, W. Electrochim. Acta 2012, 84, 223–234.

(19)

Cracknell, J. A.; McNamara, T. P.; Lowe, E. D.; Blanford, C. F. Dalton Trans. 2011, 40, 6668–6675.

(20)

Mano, N. Appl. Microbiol. Biotechnol. 2012, 96, 301–307.

(21)

Dos Santos, L.; Climent, V.; Blanford, C. F.; Armstrong, F. A. Phys. Chem. Chem. Phys. 2010, 12, 13962–13974.

(22)

Lopez, R. J.; Babanova, S.; Ulyanova, Y.; Singhal, S.; Atanassov, P. ChemElectroChem 2014, 1, 241–248.

(23)

Matanovic, I.; Babanova, S.; Chavez, M. S.; Atanassov, P. J. Phys. Chem. B 2016, 3634– 3641.


35

ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 36 of 38

(24)

Lalaoui, N.; Holzinger, M.; Le Goff, A.; Cosnier, S. Chem. - A Eur. J. 2016, 22, 1–8.

(25)

Xia, H.; Kitazumi, Y.; Shirai, O.; Kano, K. J. Electroanal. Chem. 2016, 763, 104–109.

(26)

Ramírez, P.; Mano, N.; Andreu, R.; Ruzgas, T.; Heller, A.; Gorton, L.; Shleev, S. Biochim. Biophys. Acta 2008, 1777, 1364–1369.

(27)

Murata, K.; Kajiya, K.; Nakamura, N.; Ohno, H. Energy Environ. Sci. 2009, 2, 1280– 1285.

(28)

Olejnik, P.; Pawłowska, A.; Pałys, B. Electrochim. Acta 2013, 110, 105–111.

(29)

Tominaga, M.; Ohtani, M.; Taniguchi, I. Phys. Chem. Chem. Phys. 2008, 10, 6928–6934.

(30)

Climent, V.; Zhang, J.; Friis, E. P.; Østergaard, L. H.; Ulstrup, J. J. Phys. Chem. C 2012, 116, 1232–1243.

(31)

Sugimoto, Y.; Kitazumi, Y.; Tsujimura, S.; Shirai, O.; Yamamoto, M.; Kano, K. Biosens. Bioelectron. 2015, 63, 138–144.

(32)

Singh, K.; McArdle, T.; Sullivan, P. R.; Blanford, C. F. Energy Environ. Sci. 2013, 6, 2460–2464.

(33)

Pankratov, D.; Sotres, J.; Barrantes, A.; Arnebrant, T.; Shleev, S. Langmuir 2014, 30, 2943–2951.

(34)

Kamitaka, Y.; Tsujimura, S.; Ikeda, T.; Kano, K. ELECTROCHEMISTRY 2006, 74, 642– 644.

(35)

Wang, S.; Huang, X.; Shan, X.; Foley, K. J.; Tao, N. Anal. Chem. 2010, 82, 935–941.

(36)

Boussaad, S.; Pean, J.; Tao, N. J. Anal. Chem. 2000, 72, 222–226.

(37)

Nakamoto, K.; Kurita, R.; Niwa, O. Anal. Chem. 2012, 84, 3187–3191.

(38)

Paulo, T. de F.; de Sousa, T. P.; de Abreu, D. S.; Felício, N. H.; Bernhardt, P. V; Lopes, L. G. de F.; Sousa, E. H. S.; Diógenes, I. C. N. J. Phys. Chem. B 2013, 117, 8673–8680.

(39)

Yang, L.; Gomez-Casado, A.; Young, J. F.; Nguyen, H. D.; Cabanas-Danés, J.; Huskens, J.; Brunsveld, L.; Jonkheijm, P. J. Am. Chem. Soc. 2012, 134, 19199–19206.

(40)

Jensen, U. B.; Ferapontova, E. E.; Sutherland, D. S. Langmuir 2012, 28, 11106–11114.

(41)

Lanzellotto, C.; Favero, G.; Antonelli, M. L.; Tortolini, C.; Cannistraro, S.; Coppari, E.; Mazzei, F. Biosens. Bioelectron. 2014, 55, 430–437.


36

Page 37 of 38

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ACS Catalysis

(42)

Malmqvist, M. Nature 1993, 361, 186–187.

(43)

Zubritsky, E. Anal. Chem. 2000, 72, 289–292.

(44)

Stenberg, E.; Persson, B.; Roos, H.; Urbaniczky, C. J. Colloid Interface Sci. 1991, 143, 513–526.

(45)

Mizutani, K.; Toyoda, M.; Sagara, K.; Takahashi, N.; Sato, A.; Kamitaka, Y.; Tsujimura, S.; Nakanishi, Y.; Sugiura, T.; Yamaguchi, S.; Kano, K.; Mikami, B. Acta Crystallogr. Sect. F. Struct. Biol. Cryst. Commun. 2010, 66, 765–770.

(46)

Bourdillon, C.; Bourgeois, J. P.; Thomas, D. J. Am. Chem. Soc. 1980, 102, 4231–4235.

(47)

Tsujimura, S.; Kuriyama, A.; Fujieda, N.; Kano, K.; Ikeda, T. Anal. Biochem. 2005, 337, 325–331.

(48)

Xu, F.; Berka, R. M.; Wahleithner, J. A.; Nelson, B. A.; Shuster, J. R.; Brown, S. H.; Palmer, A. E.; Solomon, E. I. Biochem. J. 1998, 334, 63–70.

(49)

Salaj-Kosla, U.; Pöller, S.; Schuhmann, W.; Shleev, S.; Magner, E. Bioelectrochemistry 2013, 91, 15–20.

(50)

Ramírez, P.; Granero, A.; Andreu, R.; Cuesta, A.; Mulder, W. H.; Calvente, J. J. Electrochem. commun. 2008, 10, 1548–1550.

(51)

McArdle, T.; McNamara, T. P.; Fei, F.; Singh, K.; Blanford, C. F. ACS Appl. Mater. Interfaces 2015, 7, 25270–25280.

(52)

Baur, J.; Le Goff, A.; Dementin, S.; Holzinger, M.; Rousset, M.; Cosnier, S. Int. J. Hydrogen Energy 2011, 36, 12096–12101.

(53)

Bryant, M. A.; Crooks, R. M. Langmuir 1993, 9, 385–387.

(54)

Lojou, E.; Bianco, P.; Bruschi, M. Electrochim. Acta 1998, 43, 2005–2013.

(55)

Buffeteau, T.; Desbat, B.; Turlet, J. M. Appl. Spectrosc. 1991, 45, 380–389.

(56)

Blaudez, D.; Turlet, J.-M.; Dufourcq, J.; Bard, D.; Buffeteau, T.; Desbat, B. J. Chem. Soc. Faraday Trans. 1996, 92, 525–530.

(57)

Rozniecka, E.; Jonsson-Niedziolka, M.; Sobczak, J. W.; Opallo, M. Electrochim. Acta 2011, 56, 8739–8745.

(58)

Matsumura, H.; Ortiz, R.; Ludwig, R.; Igarashi, K.; Samejima, M.; Gorton, L. Langmuir 2012, 28, 10925–10933.


37

ACS Catalysis

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Page 38 of 38

(59)

Schubert, K.; Goebel, G.; Lisdat, F. Electrochim. Acta 2009, 54, 3033–3038.

(60)

Gutiérrez-Sánchez, C.; Pita, M.; Vaz-Domínguez, C.; Shleev, S.; De Lacey, A. L. J. Am. Chem. Soc. 2012, 134, 17212–17220.

(61)

Baffert, C.; Sybirna, K.; Ezanno, P.; Lautier, T.; Hajj, V.; Meynial-Salles, I.; Soucaille, P.; Bottin, H.; Léger, C. Anal. Chem. 2012, 84, 7999–8005.

(62)

Alonso-Lomillo, M. A.; Rüdiger, O.; Maroto-Valiente, A.; Velez, M.; Rodríguez-Ramos, I.; Muñoz, F. J.; Fernández, V. M.; De Lacey, A. L. Nano Lett. 2007, 7, 1603–1608.

(63)

Mateo, C.; Grazu, V.; Palomo, J. M.; Lopez-Gallego, F.; Fernandez-Lafuente, R.; Guisan, J. M. Nat. Protoc. 2007, 2, 1022–1033.

(64)

Rodrigues, R. C.; Ortiz, C.; Berenguer-Murcia, Á.; Torres, R.; Fernández-Lafuente, R. Chem. Soc. Rev. 2013, 42, 6290–6307.

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