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Tryptophan Repressor-Binding Proteins from Escherichia coli and Archaeoglobus fulgidus as New Catalysts for 1,4-Dihydronicotinamide Adenine Dinucleotide-Dependent Amperometric Biosensors and Biofuel Cells Muhammad Nadeem Zafar, Federico Tasca, Lo Gorton, Eric V. Patridge, James G. Ferry, and Gilbert No#ll Anal. Chem., 2009, 81 (10), 4082-4088• DOI: 10.1021/ac900365n • Publication Date (Web): 13 April 2009 Downloaded from http://pubs.acs.org on May 14, 2009

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Anal. Chem. 2009, 81, 4082–4088

Tryptophan Repressor-Binding Proteins from Escherichia coli and Archaeoglobus fulgidus as New Catalysts for 1,4-Dihydronicotinamide Adenine Dinucleotide-Dependent Amperometric Biosensors and Biofuel Cells Muhammad Nadeem Zafar,† Federico Tasca,† Lo Gorton, Eric V. Patridge,‡ James G. Ferry,‡ and Gilbert No¨ll*,†,§ Department of Analytical Chemistry/Biochemistry, Lund University, P.O. Box 124, SE-221 00 Lund, Sweden, Department of Biochemistry and Molecular Biology, Eberly College of Science, Pennsylvania State University, 205 South Frear Laboratory, University Park, Pennsylvania, 16802-4500, and University of Siegen, Organic Chemistry 1, Adolf-Reichwein-Strasse 2, D-57068 Siegen, Germany The tryptophan (W) repressor-binding proteins (WrbA) from Echerichia coli (EcWrbA) and Archaeoglobus fulgidus (AfWrbA) were investigated for possible use in 1,4dihydronicotinamide adenine dinucleotide (NADH) dependent amperometric biosensors and biofuel cells. EcWrbA and AfWrbA are oligomeric flavoproteins binding one flavin mononucleotide (FMN) per monomer and belonging to a new family of NAD(P)H:quinone oxidoreductases (NQOs). The enzymes were covalently linked to a low potential Os redox polymer onto graphite in the presence of single-walled carbon nanotube (SWCNT) preparations of varying average lengths. The performance of the enzyme modified electrodes for NADH oxidation was strongly depending on the average length of the applied SWCNTs. By blending the Os redox polymer with SWCNTs, the electrocatalytic current could be increased up to a factor of 5. Results obtained for AfWrbA modified electrodes were better than those for EcWrbA. For NADH detection, a linear range between 5 µM and 1 mM, a lower limit of detection of 3 µM, and a sensitivity of 56.5 nA µM-1 cm-2 could be reached. Additionally spectroelectrochemical measurements were carried out in order to determine the midpoint potentials of the enzymes (-115 mV vs NHE for EcWrbA and -100 mV vs NHE for AfWrbA pH 7.0). Furthermore, an AfWrbA modified electrode was used as an anode in combination with a Pt black cathode as a biofuel cell prototype. To catalyze the oxidation of 1,4-dihydronicotinamide adenine dinucleotide (NADH) is an important task in bioanalytical chemistry, because NAD+-dependent dehydrogenases, which catalyze the oxidation of specific substrates with concomitant reduction * To whom correspondence should be addressed. Gilbert No ¨ll, University of Siegen, Organic Chemistry 1, Adolf-Reichwein-Str. 2, D-57068 Siegen, Germany. Phone: +49 (0)271 740 4360. Fax: +49 (0)271 740 3270. E-mail: noell@chemie.unisiegen.de. † Lund University. ‡ Pennsylvania State University. § University of Siegen.

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of NAD+ to NADH, constitute the largest family of redox enzymes known today.1 In principle, a variety of amperometric biosensors can be realized by employing these enzymes. The electrons gained from substrate oxidation are transferred to NAD+ and have to be detected at the electrode in terms of NADH. Unfortunately, the direct oxidation of NADH at bare electrodes requires a large overpotential and suffers from pronounced irreversibility.1-3 Therefore a second catalytic process becomes necessary. One strategy to mediate the electron transfer (ET) from NAD+-dependent enzymes to electrodes involves the covalent attachment of pyrroloquinoline quinone (PQQ) to a cystamine monolayer on gold followed by a covalent linkage of N6-(2-aminoethyl)-NAD+.2 Within this monolayer, the ET from the attached NADH to the gold surface is mediated by PQQ. NAD+-dependent dehydrogenases such as lactate dehydrogenase can be assembled on top of the NAD+ containing layer.2 An advantage of this method is that NAD+ is immobilized at the surface, and a constant level of the NAD+/ NADH redox couple in the bulk solution is not required. On the other hand, this strategy is limited to enzyme monolayers at the electrode. Alternatively, nitrocompounds immobilized on nanostructured electrodes have been shown to catalyze the oxidation of NADH at low overpotentials.4,5 Furthermore, different types of carbon nanotubes (sometimes after further modification or combination with redox mediators) were found to catalyze the oxidation of NADH quite efficiently.6-19 Another possibility is the combination of NAD+-dependent dehydroge(1) Gorton, L.; Domı´nguez, E. In Encyclopedia of Electrochemistry, Vol. 9, Biochemistry; Wilson, G. S., Ed.; Wiley-VCH: Weinheim, 2002; pp 67143. (2) Bardea, A.; Katz, E.; Bu ¨ ckmann, A. F.; Willner, I. J. Am. Chem. Soc. 1997, 119, 9114–9119. (3) Barton, S. C.; Gallaway, J.; Atanassov, P. Chem. Rev. 2004, 104, 4867– 4886. (4) Mano, N.; Kuhn, A. Biosens. Bioelectron. 2001, 16, 653–660. (5) Mano, N.; Thienpont, A.; Kuhn, A. Electrochem. Commun. 2001, 3, 585– 589. (6) Li, X.; Zhou, H.; Yu, P.; Su, L.; Ohsaka, T.; Mao, L. Electrochem. Commun. 2008, 10, 851–854. (7) Luz, R. d. C. S.; Damos, F. S.; Tanaka, A. A.; Kubota, L. T.; Gushikem, Y. Electrochim. Acta 2008, 53, 4706–4714. 10.1021/ac900365n CCC: $40.75  2009 American Chemical Society Published on Web 04/13/2009


nases with NAD(P)H:quinone oxidoreductases (NQOs) such as diaphorase, a dimeric flavin adenin dinucleotide (FAD) containing protein, which oxidize NAD(P)H with concomitant reduction of quinones or other redox mediators.20-23 On the basis of glucose dehydrogenase and diaphorase, a glucose biosensor was developed,20 and by combination of diaphorase with alcohol, aldehyde, and formate dehydrogenase, a biofuel cell capable of oxidizing methanol completely to CO2 and H2O has been presented.21 In the latter case, benzylviologen was used as a redox mediator in order to catalyze ET from diaphorase to the anode. In our previous work we have shown that diaphorase can be covalently linked to a low potential Os redox polymer, which transfers the electrons gained from NADH oxidation to the electrode.24,25 When the Os polymer was blended with oxidatively shortened and length-separated singlewalled carbon nanotubes (SWCNTs), the catalytic current for NADH oxidation could be increased by a factor of 5.24 In this contribution, we report on the performance of the tryptophan (W) repressor-binding protein (WrbA) from Echerichia coli (EcWrbA) and Archaeoglobus fulgidus (AfWrbA) with respect to applications in biosensors and biofuel cells.26-29 WrbA is an oligomeric flavoprotein that binds one flavin mononucleotide FMN per monomer. The molecular mass of monomeric EcWrbA is 21 kDa and that of AfWrbA is 22 kDa.26 EcWrbA was discovered in 1993, when it was copurified with the tryptophan repressor (TrpR).30 For a long time, the function of WrbA was not known. With respect to its redox properties, a role in oxidative stress defense was implicated.31 In line with these findings, WrbA has been recently identified as a new family of NQOs.26,28 Since for (8) Manso, J.; Mena, M. L.; Yanez-Sedeno, P.; Pingarron, J. M. Electrochim. Acta 2008, 53, 4007–4012. (9) Radoi, A.; Compagnone, D.; Valcarcel, M. A.; Placidi, P.; Materazzi, S.; Moscone, D.; Palleschi, G. Electrochim. Acta 2008, 53, 2161–2169. (10) Du, P.; Liu, S.; Wu, P.; Cai, C. Electrochim. Acta 2007, 53, 1811–1823. (11) Chakraborty, S.; Retna Raj, C. Electrochem. Commun. 2007, 9, 1323–1330. (12) Huang, M.; Jiang, H.; Qu, X.; Xu, Z.; Wang, Y.; Dong, S. Chem. Commun. 2005, 5560–5562. (13) Zhang, M.; Gorski, W. J. Am. Chem. Soc. 2005, 127, 2058–2059. (14) Valentini, F.; Salis, A.; Curulli, A.; Palleschi, G. Anal. Chem. 2004, 76, 3244– 3248. (15) Zhu, L.; Zhai, J.; Yang, R.; Tian, C.; Guo, L. Biosens. Bioelectron. 2007, 22, 2768–2773. (16) Raj, C. R.; Chakraborty, S. Biosens. Bioelectron. 2006, 22, 700–706. (17) Wang, J. Electroanalysis 2005, 17, 7–14. (18) Yan, Y.-M.; Yehezkeli, O.; Willner, I. Chem.sEur. J. 2007, 13, 10168–10175. (19) Willner, I.; Katz, E. Angew. Chem., Int. Ed. 2000, 39, 1181–1218. (20) Antiochia, R.; Gorton, L. Biosens. Bioelectron. 2007, 22, 2611–2617. (21) Palmore, G. T. R.; Bertschy, H.; Bergens, S. H.; Whitesides, G. M. J. Electroanal. Chem. 1998, 443, 155–161. (22) Gros, P.; Comtat, M. Biosens. Bioelectron. 2004, 20, 204–210. (23) Montagne, M.; Durliat, H.; Comtat, M. Anal. Chim. Acta 1993, 278, 25– 33. (24) Tasca, F.; Gorton, L.; Wagner, J. B.; No ¨ll, G. Biosens. Bioelectron. 2008, 24, 272–278. (25) Nikitina, O.; Shleev, S.; Gayda, G.; Demkiv, O.; Gonchar, M.; Gorton, L.; Csoeregi, E.; Nistor, M. Sens. Actuators, B 2007, B125, 1–9. (26) Patridge, E. V.; Ferry, J. G. J. Bacteriol. 2006, 188, 3498–3506. (27) Andrade, S. L. A.; Patridge, E. V.; Ferry, J. G.; Einsle, O. J. Bacteriol. 2007, 189, 9101–9107. (28) Carey, J.; Brynda, J.; Wolfova, J.; Grandori, R.; Gustavsson, T.; Ettrich, R.; Smatanova, I. K. Protein Sci. 2007, 16, 2301–2305. (29) Wolfova, J.; Brynda, J.; Mesters, J. R.; Carey, J.; Grandori, R.; Smatanova, I. K. Mater. Struct. Chem., Biol., Phys. Technol. 2008, 15, 55–57. (30) Yang, W.; Ni, L.; Somerville, R. L. Proc. Natl. Acad. Sci. U.S.A. 1993, 90, 5796–5800. (31) Natalello, A.; Doglia, S. M.; Carey, J.; Grandori, R. Biochemistry 2007, 46, 543–553.

EcWrbA and AfWrbA catalytic activity toward oxidation of NADH has been reported,26 it is of interest to study these enzymes with respect to applications in NAD+/NADH-dependent amperometric biosensors and biofuel cells. Similar to diaphorase, EcWrbA and AfWrbA can be covalently linked to a low potential Os redox polymer. The performance of both enzymes in the presence of different types of SWCNTs was compared. Furthermore, spectroelectrochemical measurements were carried out in order to determine the redox potentials of the enzymes. EXPERIMENTAL PART Chemicals and Materials. EcWrbA and AfWrbA were prepared as described elsewhere.26 Water was purified in a Milli-Q water purification system (Millipore, Bedford, MA). Poly(ethylene glycol) (400) diglycidyl ether (PEGDGE) was obtained from Aldrich (http:// www.sigmaaldrich.com). Poly(vinylpyridine)-[osmium-(N,N′-methylated-2,2′-biimidalzole)3]2+/3+ was synthesized as reported elsewhere.32 Single-walled carbon nanotubes (SWCNTs) were purchased from Nanocyl, Sambreville, Belgium. Triton X-100 and controlled pore glass (CPG 3000 Å) were both from Fluka (Buchs, Switzerland). Spectrographic graphite electrodes and homemade pyrolytic graphite electrodes were used. Pyrolytic graphite (PG) was obtained as a gift from Mr. Robert Pulley, Minerals Technologies (mineralstech.com). Spectrographic graphite 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 (Millipore, Bedford, MA), and the NADH used as a substrate was dissolved in 0.1 M MOPS buffer solutions. For flow injection measurements, the working buffer solutions were degassed before use to avoid air bubbles in the flow system. For voltammetric measurements (unless otherwise stated), argon was purged through the solutions for some minutes prior to the experiments. Flow injection measurements were performed with a flow-through amperometric cell of the wall-jet type33 at an applied potential of +290 mV vs NHE. The carrier flow was maintained at a constant flow rate of 1 mL min-1 by a peristaltic pump. The injection loop volume was 50 µL. The dispersion factor34 of the system was 1.04 at this flow rate. Voltammetric measurements were performed with an EG&G potentiostat/galvanostat model 273 A or an Autolab potentiostat/galvanostat PGSTAT30 (Eco Chemie, Utrecht, The Netherlands) using modified electrodes as the working electrode, a saturated calomel reference electrode (SCE), and a platinum foil counter electrode. All potentials discussed in the main part are referred to the normal hydrogen electrode (NHE). The current densities were calculated with respect to the geometric electrode area. The spectroelectrochemical setup used in this work has been described elsewhere.35 As working electrode, a gold capillary cell with an optical path length of 1 cm was applied. The extinction coefficients of the enzymes were ε450 ) 11.6 mM-1 cm-1 for EcWrbA and ε457 ) 14.0 mM-1 cm-1 for AfWrbA.26 As redox (32) Mao, F.; Mano, N.; Heller, A. J. Am. Chem. Soc. 2003, 125, 4951–4957. (33) Appelqvist, R.; Marko-Varga, G.; Gorton, L.; Torstensson, A.; Johansson, G. Anal. Chim. Acta 1985, 169, 237–247. (34) Ruzicka, J.; Hansen, E. H. In Flow Injection Analysis, 2nd ed.; Winefordner, J. D., Ed.; John Wiley & Sons: New York, 1988; pp 23-26. (35) Bistolas, N.; Christenson, A.; Ruzgas, T.; Jung, C.; Scheller, F. W.; Wollenberger, U. Biochem. Biophys. Res. Commun. 2004, 314, 810–816.

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mediators,36-38 2,6-dichloroindophenol, phenazine methosulfate, methylene blue, resazurin, 2-OH-1,4-naphthoquinone, anthraquinone 2,6-disulfonate, safranine T, diquat, 1,1′-bis(hydroxyethyl)-4,4′bipyridyl dichloride, and 1,1′-propylene-2,2′-bipyridylium dibromide were applied. Electrode Preparation and Equipment. Spectrographic graphite electrodes were polished as reported previously.39 In this work, 0.7 mg of the different SWCNT preparations was dissolved in 1 mL of Milli-Q water and sonicated overnight. Then 5 µL of dispersion was placed on the top of the polished electrode and spread evenly using a microsyringe tip. Next, 2 µL of the osmium redox polymer (10 mg/mL in Milli-Q water) was mixed with 5 µL of SWCNT solution. Following this, 5 µL of enzyme (0.1 mg/ mL in Milli-Q water) was added to the mixture. Finally 1 µL of PEGDGE (1 mg/mL) was added. The electrode was then allowed to dry and placed overnight at 4 °C in a water saturated atmosphere for the complete cross-linking reaction to occur. Electrodes not including SWCNTs were prepared as described above but without any addition of the SWCNT dispersions. Oxidative Shortening and Length Separation of SWCNTs. In order to perform the oxidative shortening, a suspension of 200 mg of SWCNTs and a mixture of 6 mL of sulfuric acid and 2 mL of nitric acid (98% and 70%, respectively) was sonicated for 12 h at 40 °C. The solution was then adjusted to pH 7 by adding a solution of NaOH. Thereafter the solvent was removed by centrifugation. To remove amorphous carbon, a suspension of the SWCNTs in Piranha solution with a 4:1 ratio of sulfuric acid (98%) and hydrogen peroxide (30 wt %) (note, this solution has to be treated with great care) was sonicated at 70 °C for 2 h. The solution was then again neutralized with NaOH and centrifuged. Next, the SWCNTs were dissolved in a 1 wt % Triton X-100 Milli-Q water solution and stabilized by sonication overnight. The dispersion was length separated by size exclusion chromatography using a column filled with controlled pore glass (CPG 3000 Å). Milli-Q water was used as the eluent. A total of 60 fractions of equal volume (15 mL) containing SWCNTs were collected. Cryogenic Transmission Electron Microscopy (CryoTEM). Specimens for electron microscopy were prepared in a controlled environment vitrification system (CEVS) to ensure stable temperature and to avoid loss of solution during sample preparation. The specimens were prepared as thin liquid films, <300 nm thick, on lacey carbon filmed copper grids and plunged into liquid ethane at -180 °C. This leads to vitrified specimens, avoiding component segmentation and rearrangement, and water crystallization, thereby preserving original microstructures. The vitrified specimens were stored under liquid nitrogen until measured. An Oxford CT3500 cryoholder and its workstation were used to transfer the specimen into the electron microscope (Philips CM120 BioTWIN Cryo, Amsterdam, The Netherlands) equipped with a postcolumn energy filter (Gatan GIF100, Warrendal, PA). The acceleration voltage was 120 kV. The images were recorded digitally with a CCD camera under low electron dose conditions. (36) Dutton, P. L. Methods Enzymol. 1978, 54, 411–435. (37) Wilson, G. S. Methods Enzymol. 1978, 54, 396–410. (38) No ¨ll, G.; Hauska, G.; Hegemann, P.; Lanzl, K.; No ¨ll, T.; von Sanden-Flohe, M.; Dick, B. ChemBioChem 2007, 8, 2256–2264. (39) Tasca, F.; Timur, S.; Ludwig, R.; Haltrich, D.; Volc, J.; Antiochia, R.; Gorton, L. Electroanalysis 2007, 19, 294–302.

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RESULTS AND DISCUSSION Flow Injection Analysis and Evaluation of SWCNTs. The enzymes EcWrbA and AfWrbA were covalently linked to a low potential Os redox polymer onto graphite electrodes and investigated with respect to their ability to catalyze the oxidation of NADH. Similar to a previous study, the Os polymer was blended with oxidatively shortened and length-separated SWCNTs in order to increase the electrocatalytic current. The SWCNTs were shortened by acid treatment for a period of 12 h and subsequently separated by size exclusion chromatography with respect to their different average lengths. A total of 60 fractions of equal volume (15 mL) were collected. For flow injection analysis (FIA), the WrbA/Os redox polymer/SWCNT coated spectrographic graphite electrodes were placed in a flow through electrochemical cell and examined in the flow injection mode. A constant potential of +290 mV vs NHE24 was applied, and the NADH oxidation current was measured at NADH concentrations of 0.5 and 1 mM (see Figure 1A). As shown in Figure 1A, the highest catalytic current was detected for AfWrbA/Os redox polymer modified electrodes blended with SWCNTs of fraction 10. The maximum current was 5 times higher than measured for electrodes prepared in the absence of SWCNTs (not shown). Similar results were obtained for EcWrbA/Os redox polymer modified electrodes. As reported previously for diaphorase/Os redox polymer modified electrodes blended with SWCNTs,24 the average length of the SWCNTs has a strong influence on the magnitude of the electrocatalytic current. When AfWrbA and EcWrbA modified electrodes were compared (see Figure 1B), the better performance (i.e., higher electrocatalytic current and higher linear current range) were found for AfWrbA. As shown in Figure 1C, for an AfWrbA/Os redox polymer/SWCNTs fraction 10 modified electrode, the linear range for NADH detection was between concentrations of 5 µM and 1 mM. The limit of detection (LOD) was at 3 µM, and a sensitivity of 56.5 nA µM-1 cm-2 was calculated. Hence, the performance of the AfWrbA/Os redox polymer/SWCNT fraction 10 modified electrode is similar to that of a diaphorase/Os redox polymer/ SWCNT modified electrode (linear range, 5 µM-7 mM; LOD, 1 µM; sensitivity, 47.4 nA µM-1 cm-2), which has been published previously.24 Cryo-Transmission Electron Microscopy. In order to obtain some information about the morphology of the SWCNTs of fraction 10, cryo-transmission electron microscopy (cryoTEM) measurements were carried out. CryoTEM was chosen as the appropriate method, because prior to conventional TEM measurements, the solvent has to evaporate, and during this process, the SWCNTs show a strong tendency to agglomerate. In Figure 2, two cryoTEM images of SWCNTs of fraction 10 are presented. As obvious from Figure 2, the average length of the carbon nanotubes is beyond 100 nm. Even though some SWCNTs are separated, the majority of SWCNTs is arranged in small bundles. Therefore the determination of the average length is not possible. However, besides the presence of ice due to freezing of the samples, the SWCNT sample seems to be of high purity. No amorphous carbon or transition metal catalyst particles, which are required for the manufacturing process of the SWCNTs, could be traced.


Figure 2. Two representative cryoTEM images of SWCNTs of fraction 10 at different magnifications

Figure 1. (A) Response of AfWrbA/Os redox polymer modified electrodes blended with different fractions of SWCNTs (fr 1, longest average length; fr 50, shortest) after injection of 0.5 and 1 mM NADH. (B) Comparison of the response of an AfWrbA and an EcWrbA/Os redox polymer modified electrode, blended with SWCNTs of fraction 10. (C) NADH calibration curves for an AfWrbA/Os redox polymer modified electrode blended with SWCNTs of fraction 10 in the concentration range from 5 µM to 10 mM NADH. Inset: Concentration range from 5 to 100 µM. Experiments were performed in 0.1 M MOPS buffer at pH 7.5.

Cyclic Voltammetry. NADH is an expensive fuel for any biofuel cell application, but it can be recovered by the oxidation of cheaper substrates such as alcohols.21 While for biosensors usually high sensitivity and substrate specificity, fast response time, and a low limit of detection are desired, for biofuel cell applications high current density and good long-term stability of the enzyme modified electrodes are required. In order to study the EcWrbA and AfWrbA/Os redox polymer/SWCNTs fraction 10 modified electrodes with respect to biofuel cell applications, cyclic voltammograms (CVs) were measured at low scan rate (v ) 0.5 mV s-1) for different concentrations of NADH and in the absence of substrate (see Figure 3). In the CVs measured in the absence of substrate, the oxidation and reduction of the Os

redox centers with a midpoint potential of about 40 mV can be seen. At an NADH concentration of 1 mM, the current density reached its maximum at potential values of less than 100 mV (at this potential, the vast majority of the Os redox centers are in their oxidized state). This observation implies that the specific analytical detection of NADH should be possible even at a potential as low as 100 mV. Nevertheless, a potential of 290 mV as applied previously24 is sufficiently low to avoid nonspecific oxidation of interfering compounds. An important difference in the performance of EcWrbA and AfWrbA becomes obvious, when the CVs measured at NADH concentrations of 10 mM are compared. For EcWrbA, the catalytic current density at 100 mV is only moderate (J ) 70 µA cm-2) but increases further when the potential is raised to more positive values (see Figure 3A). In contrast, for AfWrbA at a potential slightly below 100 mV, the catalytic current density has reached a maximum of about 240 µA cm-2 (determined by subtracting the capacitive current) and remains at this high level when the potential is further increased (see Figure 3B). A similar tendency was observed when the electrode material was changed from spectrographic graphite to pyrolytic graphite. The difference in behavior between EcWrbA and AfWrbA, as seen in parts A and B of Figure 3, could be ascribed to a suggested difference in flavin affinities. It has been noted that the flavin in EcWrbA is lost during purification while that of AfWrbA is not.26 Perhaps the behavior of EcWrbA in Figure 3A is indicative of partial cofactor dissociation.26,40 EcWrbA and (40) Ji, H.-F.; Shen, L.; Carey, J.; Grandori, R.; Zhang, H.-Y. THEOCHEM 2006, 764, 155–160.

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Figure 3. CVs of an EcWrbA/Os redox polymer/SWCNTs fraction 10 modified spectrographic graphite electrode (A) and an AfWrbA/ Os redox polymer/SWCNTs fraction 10 modified spectrographic graphite electrode (B) measured at a scan rate of v ) 0.5 mV s-1 in the presence of different concentrations of NADH and in the absence of substrate. Experiments were performed in 0.1 M MOPS buffer at pH 7.5.

AfWrbA exist as oligomers, and also structural differences between both enzymes in the oligomeric state might affect the catalytic activity. Mass spectrometry has shown that FMN promotes EcWrbA association into tetramers, which are more stable than dimers or monomers.31 High NADH concentration might have a similar or a conformational effect on EcWrbA. In order to check the long-term stability of an AfWrbA/Os redox polymer/SWCNTs fraction 10 modified electrode, multicycle CVs at a scan rate of 0.1 mV s-1 were measured over a period of 20 h. Within 20 h, the decrease in electrocatalytic current density was about 10%. Spectroelectrochemistry. In order to gain more information about the redox properties of EcWrbA and AfWrbA and to determine the midpoint potentials of both enzymes, spectroelectrochemical measurements were carried out. In Figure 4, the spectral changes during the reduction of EcWrbA and AfWrbA are shown. There is a minor experimental error (±20 mV), mainly due to the presence of redox mediators, which contribute to the absorption spectra depending on their redox states.36-38 The presence of mediators was required in order to establish electrochemical equilibrium. Within the experimental error, the same midpoint potentials were measured during reduction and reoxidation. As midpoint potential, values of -115 mV for EcWrbA and -100 mV for AfWrbA at pH 7.0 were determined as an average of three independent experiments performed at varying redox mediator concentrations. The reduction of both types of WrbA was a two electron one proton reduction without formation of a 4086

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Figure 4. Spectroelectochemical titration curves of EcWrbA (A) and AfWrbA (B) measured in the presence of a mixture of redox mediators. Depending on their redox states, the mediators are contributing to the optical spectra to some extent (the change in absorption between 550 and 700 nm is mainly caused by the mediator mixture). For both enzymes, the complete set of spectra collected during ongoing reduction is shown. The missing absorption in the spectra around 665 nm is an error of the instrument caused by the deuterium light source. Experiments were performed at 0.1 M KCl in 0.1 M MOPS buffer, pH 7.0.

stable negatively charged flavosemiquinone radical anion or a neutral flavosemiquinone radical within the time scale of the experiment.38,41 With dependence on the pKa of the doubly reduced flavohydroquinone, a second protonation step may follow the second ET leading to the flavohydroquinone in its neutral form. Any differences in the redox behavior of both enzymes, which could explain the increasing catalytic activity for EcWrbA with increasing redox potential at high NADH concentration, could not be observed. Biofuel Cell Performance. As model for a membraneless biofuel cell, an AfWrbA/Os redox polymer/SWCNTs fraction 10 modified electrode was applied as the anode together with a Pt black electrode as the cathode in a solution of NADH (8 mM) in MOPS buffer (0.1 M) at pH 7.5. Oxygen was gently purged around the cathode. Since the area of the cathode was much larger than that of the anode, the current density of this cell was limited by the anode. After an equilibration time of 500 s, a polarization curve was collected using linear sweep voltammetry (v ) 0.1 mV s-1) connecting the anode as the working, and the cathode as the reference and counter electrodes. The polarization curve and the dependence of the power density on the operating voltage are presented in parts A and B of Figure 5. The cell exhibited a maximum voltage (Vmax) of 300 mV, a maximum current (41) No ¨ll, G. J. Photochem. Photobiol., A 2008, 200, 34–38.


-150 mV simulating a variable load (see Figure 5C).42 In this period, the current density decreased about 65%. Possibly, some material was desorbed from the electrode surface due to the oxygen purging/nonquiescent conditions. Similar to other types of biofuel cell anodes,42 also for the AfWrbA/Os redox polymer/ SWCNTs fraction 10 modified electrode, the strongest decrease in activity was observed during the first hours of performance. In another stability experiment, the Pt electrode was cleaned and the NADH/buffer solution was replaced by a freshly prepared NADH/buffer solution, after the current density had decreased within the first hours of performance. This did not increase the catalytic current density. Hence, the maximum current density is limited by the AfWrbA/Os redox polymer/SWCNTs fraction 10 modified anode but not by limited stability of NADH43 leading to degradation products poisoning the Pt electrode.

Figure 5. Polarization curve (A) (measured with linear sweep voltammetry by scanning from 0 to -350 mV with a scan rate of 0.1 mV s-1 after an equilibration time of 500 s) and dependence of the power density on the operating voltage (B) for a membraneless biofuel cell consisting of an AfWrbA/Os redox polymer/SWCNTs fraction 10 modified electrode (basal oriented) as the anode and a Pt black electrode as the cathode. As fuel, a solution of NADH (8 mM) in MOPS buffer (0.1 M, pH 7.5) was used. For stability measurements, multicycle CVs (39 cycles) were collected at a scan rate of v ) 0.1 mV s-1 in the potential range between -50 and -150 mV in order to simulate a variable load.

density (J max) of 105 µA cm-2, and a maximum power density (Pmax) of 12 µW cm-2 at an operating voltage of 165 mV (under oxygen purging/nonquiescent conditions). A fill factor of 0.38 was calculated by dividing Pmax by the product of Vmax and Jmax. The fill factor is the ratio of the experimentally determined maximum power density divided by the maximum power density, which can be reached theoretically for a biofuel cell with a given Vmax and Jmax under ideal conditions. In order to prove the stability, the biofuel cell was investigated for a period of 21 h and 40 min by multicycle CVs at low scan rate (v ) 0.1 mV s-1) in a potential range between -50 and

CONCLUSIONS In this work, the NQOs EcWrbA and AfWrbA were investigated with respect to applications in biosensors and biofuel cells. For this purpose, the enzymes were covalently linked to a low potential Os redox polymer in the presence of oxidatively shortened and length-separated SWCNTs. By blending the Os redox polymer with SWCNTs, the electrocatalytic current could be increased up to a factor of 5. In line with previous studies on the NQO diaphorase,24 in the current study a dependence of the electrocatalytic current on the average length of the SWCNTs was also found. The highest current was obtained with SWCNTs of fraction 10. When AfWrbA and EcWrbA/Os redox polymer/SWCNT fraction 10 modified electrodes were compared, the better results were obtained with AfWrbA. Also during cyclic voltammetry experiments, a higher catalytic current could be obtained, and the maximum current was reached at lower potential for AfWrbA than for EcWrbA. The AfWrbA based NADH biosensor exhibited excellent performance. The linear range for NADH detection by an AfWrbA/Os redox polymer/SWCNT fraction 10 modified electrode was between concentrations of 5 µM and 1 mM, the LOD was at 3 µM, and a sensitivity of 56.5 nA µM-1 cm-2 was calculated. These parameters are in the same range as reported previously for other types of enzymatic NADH biosensors.24 For a diaphorase/Os redox polymer/SWCNT modified electrode (linear range, 5 µM-7 mM), a slightly lower LOD (1 µM) but also a somewhat lower sensitivity (47.4 nA µM-1 cm-2) has been reported.24 Hence, AfWrbA turned out to be equally suited for the development of an NADH sensor as diaphorase (from Bacillus stearothermophilus). Possibly, future types of WrbA may be even more suited to catalyze the electrochemical oxidation of NADH than AfWrbA. In order to study EcWrbA and AfWrbA more in detail, spectroelectrochemical measurements were performed. Within the experimental error, the same midpoint potentials at pH 7.0 (-115 mV for EcWrbA and -100 mV for AfWrbA) were measured during reduction and reoxidation. For both types of WrbA, a two electron one proton reduction without formation of a stable negatively charged flavosemiquinone radical anion or a neutral flavosemi(42) Tasca, F.; Gorton, L.; Harreither, W.; Haltrich, D.; Ludwig, R.; No ¨ll, G. J. Phys. Chem. C 2008, 112, 13668–13673. (43) Chenault, H. K.; Whitesides, G. M. Appl. Biochem. Biotechnol. 1987, 14, 147–197.

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quinone radical was observed. Furthermore, an AfWrbA/Os redox polymer/SWCNTs fraction 10 modified electrode was applied as the anode together with a Pt black electrode as the cathode as a model for a membraneless biofuel cell using NADH as the substrate. This cell exhibited a maximum voltage of 300 mV, a maximum current density of 105 µA · cm-2, and a maximum power density of 12 µW cm-2 at an operating voltage of 165 mV (under oxygen purging/nonquiescent conditions). In contrast to multicycle CV experiments (with a decrease of about 10% in catalytic current density within 20 h), there was a stronger decrease in activity (65% of decrease in 21 h and 40 min) during the biofuel cell experiments. Possibly, this was due to limited mechanical stability of the electrocatalytically active layer at the electrode surface under oxygen purging/ nonquiescent conditions.

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ACKNOWLEDGMENT M.N.Z. and F.T. contributed equally to this work. 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 (Projects 6212004-4476 and 621-2007-4124), and the “Higher Education CommissionofPakistan”.WethankGunnelKarlssonattheBiomicroscopy Unit, Polymer and Materials Chemistry, Institute of Chemistry, Lund University, Lund, Sweden, for performing the cryoTEM work.

Received for review February 17, 2009. Accepted March 24, 2009. AC900365N

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