Direct Electron Transfer at Cellobiose Dehydrogenase Modified ...

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J. Phys. Chem. C 2008, 112, 9956–9961

Direct Electron Transfer at Cellobiose Dehydrogenase Modified Anodes for Biofuel Cells Federico Tasca,† Lo Gorton,† Wolfgang Harreither,‡ Dietmar Haltrich,‡ Roland Ludwig,§ and Gilbert No¨ll*,† Department of Analytical Chemistry, Lund UniVersity, P.O. Box 124, SE-221 00 Lund, Sweden, DiVision of Food Biotechnology, Department of Food Sciences and Technology, BOKU-UniVersity of Natural Recources and Applied Life Sciences Vienna, Muthgasse 18, A-1190 Wien, Austria, and Research Centre Applied Biocatalysis, Petersgasse 14, A-8010 Graz, Austria ReceiVed: March 10, 2008; ReVised Manuscript ReceiVed: April 17, 2008

Cellobiose dehydrogenases (CDHs, EC 1.1.99.18) contain a larger flavin-associated (dehydrogenase) domain and a smaller heme-binding (cytochrome) domain. CDHs from basidiomycete fungi oxidize at an appreciable level cellobiose, cellodextrins, and lactose, and those from ascomycetes may additionally oxidize some monosaccharides to their corresponding lactones at the flavin domain. CDHs are able to communicate directly with an electrode via their heme domain. In this work, different types of CDHs have been adsorbed on graphite electrodes and studied with respect to their direct electron transfer (DET) properties. Electrochemical studies were performed in the presence and absence of single-walled carbon nanotubes (SWCNTs) using lactose as substrate. In the presence of SWCNTs, the electrocatalytic current for substrate oxidation based on DET between enzyme and electrode was significantly increased. Furthermore, the onset of the electrocatalytic current was at lower potential than in the absence of SWCNTs. The highest electrocatalytic activity toward oxidation of lactose was found for CDH from the basidiomycete Phanerochaete sordida. Based on CDH from Phanerochaete sordida, an anode for biofuel cells was developed. This anode using lactose as substrate was combined with a Pt black cathode for oxygen reduction as a model for a membrane-less biofuel cell in which the processes at both electrodes occur by DET. Introduction Redox enzymes catalyze the oxidation or reduction of a specific substrate or a group of substrates usually with similar structural and electronic properties. Depending on their enzymatic function these enzymes can be applied in biosensors1–3 or biofuel cells.4–7 While for biosensor applications, high sensitivity and substrate selectivity have to be reached, biofuel cells require modified electrodes working with high current densities. At the same time, long-term stability is desired. This is in contrast to biosensors which have to be cheap and are often designed for fast response time and single use. For both applications, the electrochemical addressability of the redox enzymes has to be achieved. This is possible by the use of redox mediators.8,9 When enzymes are “wired” to osmium redox polymers, the electron transfer (ET) is mediated by flexible Os2+/3+ redox centers which are able to transport the charge from the electrochemically active part of the enzyme to the electrode.7,10 As an alternative to mediated electron transfer (MET), some enzymes are able to communicate directly with the electrode. The waiving of redox mediators in biofuel cell or biosensor applications simplifies the electrode architecture and reduces the amount of components which have to be optimized. Furthermore, the maximum cell voltage of biofuel cells is usually decreased by the use of redox mediators (for thermodynamic reasons).7 Currently, direct electron transfer * Corresponding author. Phone: +46 46 222 0103. Fax: +46 46 222 4544. E-mail: [email protected]. New address: Max Planck Institute for Polymer Research, Ackermannweg 10, 55128 Mainz, Germany † Lund University. ‡ BOKU-University of Natural Recources and Applied Life Sciences Vienna. § Reseach Centre Applied Biocatalysis.

(DET) has been reported only for about 50 out of 1060 redox enzymes know today.2,11–13 In recent years, the sensitivity of amperometric biosensors consisting of enzymes embedded in osmium redox polymers could be improved by the introduction of carbon nanotubes.14–16 During investigations of redox enzymes with respect to biosensor or biofuel cell applications, it also turned out that the rate of DET can be increased by single-walled17–20 or multiwalled21–29 carbon nanotubes. In this contribution, the DET properties of different types of cellobiose dehydrogenase (CDHs) in the presence and absence of singlewalled carbon nanotubes (SWCNTs) are compared with respect to potential applications in biofuel cells. CDHs have been used previously for the development of amperometric biosensors based on DET.30–33 CDHs are extracellular enzymes produced by a variety of fungi from the phyla Basidiomycota and Ascomycota.13,34 The enzymes exist usually as monomers containing a larger flavin-associated (dehydrogenase) domain and a smaller heme containing (cytochrome) domain.34 The cofactors in the flavin and heme domain are flavin adenine dinucleotide (FAD) and heme b.34 CDHs oxidize cellobiose, cellodextrins, and lactose at the flavin domain.30–34 When CDHs are immobilized on electrodes, oxidation of a substrate at the flavin domain is followed by intramolecular ET to the heme domain, which is able to communicate with the electrode via DET.13,30–33 Alternatively, the electrons can be transferred to the electrode from either the flavin or the heme domain by MET.13,30 Whereas MET has been shown for the intact enzyme as well as for the separated flavin domain, efficient DET requires the presence of the heme domain.13,30,35 We have screened five different types of CDH with respect to their DET properties after coadsorption with SWCNTs on graphite electrodes. The CDHs from the basidiomycete fungi Phanerochaete sordida31,32

10.1021/jp802099p CCC: $40.75  2008 American Chemical Society Published on Web 06/11/2008

DET in a CDH Biofuel Cell

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TABLE 1: Molecular Properties and Kinetic Constants of the Investigated CDHs (EC 1.1.99.18) and Activities of the Samples Which Have Been Used CDH Ps Pc Tv43 Mt Sr42

Holo-enzyme/Da 88 9044 89 94 101

glycosylation [a]/%

pH optimum DCIP

KM lactose/mM

kcat lactose/s-1

Sample activity/U · ml-1

Specific activity/U · mg-1

[b

4.0 5.045 4.5 6.0 4.0

2.3 1.145 3.7 0.06 2.4

14.0 13.445 26.9 17.2 26.0

140 50 140 360 210

33 39 69 10 63

] 10 11 8 15/20 [a]

a The percentage of glycosylation was approximated by subtracting the molecular weight of the available protein sequences from the total molecular weights obtained by SDS-PAGE, gel filtration, or MALDI-MS. b Amino acid sequence not known.

(PsCDH), Phanerochaete chrysosporium33 (PcCDH), and Trametes Villosa31–33(TVCDH), from the ascomycete fungus Myriococcum thermophilum30,31 (MtCDH), and from the plant pathogen fungus Sclerotium rolfsii36 (SrCDH) were compared. Experimental Section Water was purified in a Milli-Q water purification system (Millipore, Bedford, MA). Triton X-100 and controlled pore glass (CPG 3000 Å) were both from Fluka (Buchs, Switzerland). Poly(ethylene glycol) (400) diglycidyl ether (PEGDGE) was obtained from Aldrich (http://www.sigmaaldrich.com). Single-walled carbon nanotubes were purchased from Nanocyl, Sambreville, Belgium. Spectrographic graphite rod electrodes were from Ringsdorff Werke GmbH, Bonn, Germany, (type RW001, 3.05 mm diameter and 13% porosity http://www.sglcarbon.com/). All solutions used for immobilization were prepared in Milli-Q water. Electrode Preparation and Equipment. Flow injection measurements were performed with a flow-through amperometric cell of the wall-jet type.37 The carrier flow was maintained at a constant flow rate of 0.5 mL min-1 by a peristaltic pump. The injection loop volume was 50 µL. The dispersion factor38 of the system was 1.04 at this flow rate. These experiments were performed in the presence of oxygen. Cyclic voltammetry was performed with an EG&G potentiostat/ galvanostat model 273A using modified electrodes as working electrode, a saturated calomel reference electrode (SCE), and a platinum foil counter electrode. Before measuring a cyclic voltammogram, argon was purged through the solutions for some minutes. All potentials discussed in the main part are referred to the normal hydrogen electrode (NHE). Graphite rod electrodes were polished as reported previously.39 From each nanotube preparation, 0.7 mg were sonicated in 1 mL of Milli-Q water prior to use. A 5-10 µL sample of the nanotube suspension was placed on the polished end of the electrode and spread evenly using the microsyringe tip. Next, 5-10 µL of enzyme solution was mixed to the previously added suspension. Optionally, 1-2 µL of poly(ethylene glycol) diglycidyl ether (PEGDGE; 2.5-20 mg/ml) was added. The electrodes were then allowed to dry and placed overnight at 4°C for complete cross-linking reaction. Electrodes not containing SWCNTs were prepared as described above without adding a nanotube suspension. Oxidative Shortening and Length Separation of the SWCNTs.40 The oxidative shortening and length separation were performed as described elsewhere.41 Preparation and Characterization of the Enzymes. SrCDH was produced from cultures of Sclerotium rolfsii CBS 191.62 and purified as previously described.42 MtCDH was obtained from Myriococcum thermophilum CBS 208.89 using a published protocol,30 and TVCDH from Trametes Villosa CBS 334.49 was produced according the method from Ludwig et al.43 The same

procedure for CDH production and purification was used to obtain PcCDH from Phanerochaete chrysosporium K3 (a kind gift of Dr. Jindrich Volc, Institute of Microbiology, Academy of Sciences of the Czech Republic) and PsCDH from Phanerochaete sordida MB 66 (a kind gift of Dr. Hansjo¨rg Prillinger, Institute of Applied Microbiology, University of Natural Resources and Applied Life Sciences, Vienna). Molecular properties and kinetic parameters of the studied enzymes are summarized in Table 1. Most data are collected from published sources and indicated by a reference. Other data were determined using published methods.42 The maximum electrocatalytic current for oxidation of 10 mM lactose was almost constant, when the amount of enzyme solution used for electrode preparation was varied between 5 or 10 µL or when samples of different activities were compared for the same enzyme indicating that an excess of enzyme was present. Results and Discussion DET Properties of the CDHs. We first investigated MtCDH which was recently discovered from an ascomycete fungus, because for this CDH the ability to oxidize a broad range of substrates was reported.30 The enzyme was adsorbed to graphite electrodes in the presence and absence of SWCNTs. Besides unmodified SWCNTs, also oxidatively shortened SWCNTs40 purified and fractionated by size exclusion chromatography were examined. In a typical procedure, 5 µL of enzyme solution was adsorbed on a graphite electrode in the presence or absence of 5 µL of the nanotube suspension (0.7 mg ml-1). The electrodes were compared with respect to their ability to oxidize lactose at concentrations of 10, 25, and 50 µM. Lactose was chosen as substrate, because all CDHs exhibit high activity for this substrate, and in contrast to the cellodextrins, there is no substrate inhibition.30,33 The modified electrodes were placed in a flow through electrochemical cell and examined in the flow injection mode at a constant applied potential (+588 mV vs the normal hydrogen electrode, NHE) for their response to injected samples of lactose solutions. The highest increase in catalytic current was measured in the presence of unmodified SWCNTs. Since it was our main interest to develop an anode for biofuel cell applications, we improved the electrode modification protocol with unmodified SWCNTs, which had shown the highest electrocatalytic current. The electrodes became more stable by adding a solution of cross-linking reagent, poly(ethylene glycol) diglycidyl ether (PEGDGE), during adsorption of enzyme and nanotubes.46 The amount of adsorbed enzyme and nanotubes could also be doubled by this procedure. When electrodes, prepared with the same amount of SWCNTs and enzyme but in the absence of cross-linker, were studied, the electrocatalytic current strongly decreased in the first 10 min indicating that SWCNTs and enzyme are partially washed off the electrode.

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Figure 1. Polarization curves of CDH/SWCNT/cross-linker modified electrodes. The modified electrodes were placed in a flow through electrochemical cell with a 10 mM solution of lactose added to the flow buffer (0.1 M citrate buffer, pH 5.3). PsCDH, PcCDH, TVCDH, MtCDH, and SrCDH were investigated.

Next, we studied PsCDH because excellent DET properties have been reported for this CDH.31,32 When the response of these two CDHs to lactose oxidation was studied in the flow injection mode, the current could be increased up to 1 order of magnitude in the presence of SWCNTs. A calibration curve for lactose oxidation by MtCDH and PsCDH in the absence of SWCNTs led to the same results as published previously.30,32 When these enzymes were studied in the presence of SWCNTs, a calibration curve exhibiting a linear range between current and lactose concentration could not be determined because of mass transfer resistances in the system. For a comparative study, electrodes modified with five different CDHs in the presence of SWCNTs were placed in the flow through electrochemical cell and investigated for their response to lactose using a 10 mM lactose solution at pH 5.3 (citrate buffer, 0.1 M) as the mobile phase. At a concentration of 10 mM lactose, substrate saturation is almost reached (the Michaelis-Menten constants (KM) of the different CDHs are depicted in Table 1 in the Experimental Section). A pH value of 5.3 was chosen because for this pH the highest current toward oxidation of lactose has been reported for MtCDH.30 By changing the applied potential, polarization curves were measured, which are presented in Figure 1. In line with previous investigations,31,32 PsCDH exhibits a higher rate of DET than the other CDHs. Furthermore, the electrocatalytic current starts at a lower potential than observed for the other CDHs. PcCDH, TVCDH, and MtCDH exhibit a similar response current profile (in the range between 300 and 600 mV). In a previous study, higher rates of DET have been reported for TVCDH in comparison to PcCDH.33 However, these studies were performed at different pHs and with a lactose concentration of only 200 µM which is below substrate saturation (in contrast to the lactose concentration of 10 mM applied in this work). In comparison with PsCDH, the DET rate of MtCDH was only moderate. This is in agreement with previous studies of both enzymes entrapped at thiol modified gold electrodes.31 The electrocatalytic current of MtCDH starts at the second lowest potential followed by PcCDH and TVCDH. The lowest catalytic activity and the most positive onset of the electrocatalytic current were detected for SrCDH, which is the largest and most highly glycosylated CDH (see Table 1, Experimental Section). Next, PsCDH (because of its high rate of DET) and MtCDH (due to its broad range of substrates)30 were studied in detail. For these two species also, polarization curves in the absence of SWCNTs

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Figure 2. Polarization curves for PsCDH and MtCDH modified electrodes in the presence (filled symbols) and absence (empty symbols) of SWCNTs and cross-linker. The modified electrodes were placed in a flow through electrochemical cell with a 10 mM solution of lactose added to the flow buffer (0.1 M citrate buffer, pH 5.3).

were collected as described above. In Figure 2, the polarization curves of PsCDH and MtCDH collected in the presence and absence of SWCNTs are presented. As obvious from Figure 2, coadsorption of SWCNTs results not only in an increase in the electrocatalytic current but also in a decrease in the overpotential for substrate oxidation. Cyclic and Square Wave Voltammetry. PsCDH and MtCDH were studied by cyclic and square wave voltammetry in the absence of substrate. A pH value of 4.0 for PsCDH and 5.5 for MtCDH was chosen in order to improve the conditions for DET with respect to the individual CDHs.30,32 In line with previous investigations,30,32 no well-defined electrochemical response was observed, when the enzymes were adsorbed on graphite electrodes in the absence of SWCNTs. In the presence of SWCNTs, an anodic and cathodic wave for reduction and reoxidation of CDH could be observed. Better resolution was obtained for oxidatively shortened SWCNTs compared with unmodified SWCNTs. For these investigations, 5 µL of enzyme solution was adsorbed on a graphite electrode together with 5 µL of the nanotube suspension. Representatively, one cyclic voltammogram (CV) and one square wave voltammogram for PsCDH and MtCDH are depicted in Figures 3 and 4. As obvious from Figures 3A and 4A, in the CVs, the maxima of the anodic waves are better resolved than those for the cathodic waves. In the CVs and square wave voltammograms, two electrochemical processes can be distinguished. As implied by the current ratio, the first (more positive) wave might be the reduction/reoxidation of the heme (1 e-), and the second wave might be that of the flavin domain (2 e-). From Figures 3B and 4B, the potential difference is expected to be less than 100 mV for PsCDH and about 150 mV for MtCDH. During additional experiments with PsCDH modified electrodes in the presence of 0.1 M lactose, the onset of the electrocatalytic current started with the onset of the anodic wave, which was at more positive potential. Hence, the more positive redox process could be the reduction/reoxidation of the heme domain as observed previously for CDHs immobilized at modified gold electrodes.13,31,32,36 We could not prove that there already was a significant amount of electrocatalytic current, when the more negative redox center (possibly the flavin domain) was oxidized. However, the electrodes prepared in the absence of the crosslinker were not very stable. In control experiments, we studied also the CDH modified electrodes after additional adsorption of free FAD. The reduction of free FAD was detected at more negative potentials (about 100 mV) than the second reduction depicted in Figures 3 and 4. The second (more negative) redox

DET in a CDH Biofuel Cell

Figure 3. (A) A CV of a graphite electrode modified with PsCDH + oxidatively shortened SWCNTs (fraction 3), scanned from 544 mV to -226 mV and back, V ) 0.5 mV s-1. (B) A square wave voltammogram of the same electrode; frequency, 1 Hz; pulse height, 20 mV; step, 2 mV. The experiments were carried out in a 0.1 M citrate buffer solution, pH 4.0

process might be caused by the flavin domain, which is in a catalytically inactive state, for example, due to partial denaturation during enzyme adsorption. A similar electrochemical process has been observed for PsCDH entrapped at thiol modified gold electrodes.31 If the redox processes observed for PsCDH and MtCDH (presented in Figures 3 and 4) are caused by the flavin and the heme domain, they are found at similar potentials as those observed for PcCDH. For PcCDH, the redox potentials of the flavin (106 mV, pH 3; -132 mV, pH 7) and the heme domain (190 mV, pH 3; 130 mV, pH 7) have been reported47 (according to Nernst, a shift of -59 mV per pH unit is expected for the flavin redox transformation when two protons and two electrons are involved). Also, the redox potential of the heme domain is pH dependent because some residues close to the heme change their protonation state because of different pKa values in the reduced and oxidized forms of CDH.36 Examination of CDH/SWCNT Modified Anodes with Respect to Biofuel Applications. In order to determine the electrocatalytic current under diffusion controlled conditions, we investigated the CDH/SWCNT modified electrodes in the presence of substrate by cyclic voltammetry at low scan rate (V e 1 mV s-1). A pH of 4.5 was chosen, because at this pH CDH modified anodes could be applied in membrane-less biofuel cells based on DET in combination with laccase modified cathodes.4,7 For comparison between PsCDH and MtCDH modified electrodes, a 5 mM lactose solution was chosen as substrate. In line with the measurements performed in the flow through electrochemical cell (see Figure 2), the electrocatalytic current for PsCDH was three to four times higher than that for MtCDH. Thus, in further experiments, we focused on PsCDH as enzyme for the development of biofuel cell anodes. The CVs of a

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Figure 4. (A) A CV of a graphite electrode modified with MtCDH + oxidatively shortened SWCNTs (fraction 3), V ) 50 mV s-1. (B) A square wave voltammogram of the same electrode: frequency, 5 Hz; pulse height, 50 mV; step, 5 mV. The experiments were carried out in a 0.1 M citrate buffer solution, pH 5.5. The arrow in (A) and (B) indicates a shoulder belonging to the first reduction.

Figure 5. CVs of a graphite electrode modified with PsCDH, SWCNTs, and cross-linker. The CVs were measured in buffer solution (0.1 M citrate buffer, pH 4.5) at a scan rate of V ) 1 mV s-1 in the presence of 100 mM lactose (solid line), 5 mM lactose (dotted line), 5 mM cellobiose (dashed line), and without substrate (dashed-dotted line).

PsCDH modified electrode in the presence and absence of substrate are presented in Figure 5. At the applied scan rate of 1 mV s-1, there is still some capacitive current due to charging/discharging of the electrode. In order to determine polarization curves, the forward scan in the absence of substrate was subtracted from the forward scan in the presence of substrate. To obtain the current density, the resulting current was divided by the macroscopic area of the electrode (A ) 0.0706 cm2). The polarization curves for a graphite electrode modified with PsCDH, SWCNTs, and crosslinker are depicted in Figure 6A. At a substrate concentration of 5 mM, the catalytic current starts at 100 mV and increases

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Figure 6. Polarization curves for a graphite electrode at pH 4.5 (0.1 M citrate buffer). (A) Electrodes modified with PsCDH, SWCNTs, and cross-linker. Polarization curves were measured in the presence of 100 mM lactose (solid line), 5 mM lactose (dotted line), and 5 mM cellobiose (dashed line). (B) Electrodes modified with SWCNTs and cross-linker. Polarization curves were measured in the presence of 100 mM lactose (solid line) and 5 mM lactose (dotted line).

up to a value of 250 mV. At more positive potentials, the current density remains almost constant. The highest current density was obtained for lactose as substrate. At a substrate concentration of 100 mM lactose, the current density is higher than that for 5 mM lactose and continues to increase at more positive potentials. This behavior implies that some additional noncatalytic current due to direct oxidation of lactose at the graphite, the SWCNTs, or both is present. To prove this assumption, SWCNTs were adsorbed to a graphite electrode in the absence of CDH and investigated at different lactose concentrations. As depicted in Figure 6B, there is some current density caused by nonspecific oxidation at high substrate concentration. The experiments were carried out at least three times leading to similar results. However, there was some variation in the absolute amount of current since the suspension of carbon nanotubes, which had been used for electrode modification, was not completely homogeneous, even after extended sonication. Therefore, the amount of SWCNTs adsorbed to individual electrodes may vary. The stability of the electrodes was proven by cyclic voltammetry (multicycle experiment). A graphite electrode modified with PsCDH, SWCNTs, and cross-linker was cycled at low scan rate (V ) 0.1 mV s-1) between 294 and 394 mV. After 12 h, a decrease in current of about 20% was observed. In order to study the electrode performance in the presence of oxygen a PsCDH/SWCNT/cross-linker modified electrode was applied as anode together with a Pt black cathode in a 5 mM lactose solution (at pH 4.5, citrate buffer, 0.1 M). Oxygen was gently purged through the solution close to the cathode. Since the area of the Pt cathode was much larger than that of

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Figure 7. (A) Polarization curve (measured with linear sweep voltammetry) and (B) dependence of the power density on the operating voltage for a membrane-less biofuel cell consisting of a graphite electrode modified with PsCDH, SWCNTs, and cross-linker as anode and a Pt black electrode as cathode. As fuel, a 5 mM lactose solution (0.1 M citrate buffer, pH 4.5) was used.

the anode, the current density of this cell was limited by the anode. After operating for about 1 h, a polarization curve was measured by linear sweep voltammetry (V ) 0.2 mV s-1) connecting the anode as working and the cathode as reference and counter electrode. The polarization curve and the dependence of the power density on the operating voltage are presented in Figure 7. This cell exhibited a maximum voltage of 590 mV, a maximum current density of 112 µA · cm-2, and a maximum power density of 32 µW · cm-2 at an operating voltage of 430 mV (under oxygen purging/nonquiescent conditions). A fill factor of 0.48 was calculated. By replacing the Pt cathode by a laccase or bilirubin oxidase modified cathode, the maximum voltage of the cell is expected to be further increased.7 Conclusions The coadsorption of SWCNTs with CDHs increases the DET rate and reduces the overpotential for substrate oxidation. Depending on the nature of the SWCNTs, the resolution during analytical measurements such as cyclic voltammetry can also be improved. Among the five CDHs investigated, the highest rate of DET was observed for PsCDH. A CDH modified anode has been developed, which can be applied in membrane-less biofuel cells.4,5,7 The high catalytic activity toward lactose and cellobiose might be interesting for applications in context with the production of lactose free milk, lactose measurements in the dairy industry, or exploiting cellobiose containing waste from paper industry. Furthermore, the catalytic activity of CDH toward cellobiose might be important for the future use of alternative fuels based on cellulose. The current density of the PsCDH/SWCNT/cross-linker modified anode working by DET at a lactose concentration of 5 mM is about 80 µA · cm-2. This

DET in a CDH Biofuel Cell value is the same order of magnitude as reported for anodes working by MET using glucose as fuel.18,19 These anodes were based on NAD dependent glucose dehydrogenase, poly(brilliant cresyl blue), or poly(methylene blue), and SWCNTs, and required NADH redox cycling.18,19 In a first experiment, a PsCDH/SWCNT/cross-linker modified anode was applied together with a Pt black cathode in a lactose solution as a model for a membrane-less biofuel cell, in which the processes at both electrodes occur by DET. Additional improvement of the electrode architecture will be carried out in order to increase stability and power density of the anode. Our further interest will be the construction of membrane-less biofuel cells based on DET, in which also oxygen reduction is catalyzed by redox enzymes such as laccase18,48,49 or bilirubin oxidase.19,24,50 Acknowledgment. This work was supported by the Deutsche Forschungsgemeinschaft DFG (DFG Postdoctoral Fellowship NO 740/1-1 and Ru¨ckkehrstipendium NO 740/3-1), the Swedish Research Council (Project Nos. 621-2004-4476 and 621-2007-4124) and the Austrian Science Fund (Project No. L395-B11). References and Notes (1) Habermu¨ller, K.; Mosbach, M.; Schuhmann, W. Fresenius’ J. Anal. Chem. 2000, 366, 560. (2) Wollenberger, U. Third generation biosensors - integrating recognition and transduction in electrochemical sensors. In Biosensors and Modern Biospecific Analytical Techniques; Gorton, L., Ed.; Elsevier: Amsterdam, 2005; pp 65. (3) Schuhmann, W. ReV. Mol. Biotechnol. 2002, 82, 425. (4) Barton, S. C.; Gallaway, J.; Atanassov, P. Chem. ReV. 2004, 104, 4867. (5) Bullen, R. A.; Arnot, T. C.; Lakeman, J. B.; Walsh, F. C Biosens. Bioelectron 2006, 21, 2015. (6) Willner, I.; Katz, E. Angew. Chem., Int. Ed. 2000, 39, 1181. (7) Heller, A. Phys. Chem. Chem. Phys. 2004, 6, 209. (8) Dutton, P. L. Methods Enzymol. 1978, 54, 411. (9) Wilson, G. S. Methods Enzymol. 1978, 54, 396. (10) Heller, A. Curr. Opin. Chem. Biol. 2006, 10, 664. (11) Borgmann, S.; Hartwich, G.; Schulte, A.; Schuhmann, W. Amperometric enzyme sensors based on direct and mediated electron transfer. In Electrochemistry of Nucleic Acids and Proteins; Palacek, E., Scheller, F. W., Wang, J., Eds.; Elsevier: Amsterdam, 2005; pp 599. (12) Ferapontova, E. E.; Shleev, S.; Ruzgas, T.; Stoica, L.; Christenson, A.; Tkac, J.; Yaropolov, A. I.; Gorton, L. Direct electrochemistry of proteins and enzymes. In Electrochemistry of Nucleic Acids and Proteins; Palacek, E., Scheller, F. W., Wang, J., Eds.; Elsevier: Amsterdam, 2005; pp 517. (13) Christenson, A.; Dimcheva, N.; Ferapontova, E. E.; Gorton, L.; Ruzgas, T.; Stoica, L.; Shleev, S.; Yaropolov, A. I.; Haltrich, D.; Thorneley, R. N. F.; Aust, S. D. Electroanalysis 2004, 16, 1074. (14) Antiochia, R.; Gorton, L. Biosens. Bioelectron. 2007, 22, 2611. (15) Joshi, P. P.; Merchant, S. A.; Wang, Y.; Schmidtke, D. W. Anal. Chem. 2005, 77, 3183. (16) Wang, Y.; Joshi, P. P.; Hobbs, K. L.; Johnson, M. B.; Schmidtke, D. W. Langmuir 2006, 22, 9776. (17) Zhu, L.; Yang, R.; Zhai, J.; Tian, C. Biosens. Bioelectron. 2007, 23, 528.

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