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Direct heterogeneous electron transfer reactions and molecular orientation of fructose dehydrogenase adsorbed onto pyrolytic graphite electrodes
Available online at www.sciencedirect.com Journal of
Electroanalytical Chemistry Journal of Electroanalytical Chemistry 610 (2007) 1–8 www.elsevier.com/locate/jelechem
Direct heterogeneous electron transfer reactions and molecular orientation of fructose dehydrogenase adsorbed onto pyrolytic graphite electrodes Masato Tominaga *, Chiharu Shirakihara, Isao Taniguchi
*
Graduate School of Science and Technology, Kumamoto University, Kumamoto 860-8555, Japan Received 25 April 2007; received in revised form 7 June 2007; accepted 19 June 2007 Available online 28 June 2007
Abstract Direct heterogeneous electron transfer reactions and the molecular orientation of D-fructose dehydrogenase (FDH, from Gluconobacter sp.) adsorbed onto electrodes were investigated. Catalytic oxidation currents based on the direct electron transfer reactions of FDH adsorbed onto basal-plane, highly oriented pyrolytic graphite (basal-plane HOPG), plastic formed carbon plate and tin-doped indium oxide electrodes were observed from a potential around 0.1 V (vs. Ag/AgCl/saturated KCl) in phosphate solution (pH 5.0) in the presence of fructose as a substrate for FDH. The catalytic current for the FDH adsorbed onto basal-plane HOPG electrodes indicated a pH dependence. The catalytic oxidation currents were only observed in acidic solutions (pH 6 6), and were not observed in neutral and alkaline solutions. The reason why the catalytic current was not obtained in neutral and alkaline solutions was because the FDH complex decomposed in neutral and alkaline solutions based on the AFM measurements. The differential pulse voltammetric investigations for FDH adsorbed onto basal-plane HOPG electrode together with the AFM and electrophoresis results indicated that the flavin-containing subunit in FDH accepts electrons from D-fructose, and transfers these electrons to the heme c-containing subunit, and then the direct electron transfer reaction of FDH occurred at the heme c site. Ó 2007 Published by Elsevier B.V. Keywords: Fructose dehydrogenase; Electron transfer reaction; Orientation; Catalytic current; Flavin; Heme
1. Introduction Direct heterogeneous electron transfer reactions between proteins and electrodes have been extensively studied from the viewpoints of both understanding the fundamental features of proteins, and for applications as biodevices, biosensors and enzyme-biofuel cell. The first direct electron transfer reactions of proteins were performed by the Hill and Eddowes [1], Kuwana and Yeh [2] and Niki et al. [3] groups, in 1977, 1977 and 1979, respectively, in which the reversible cyclic voltammetric responses of cytochrome c on gold and tin-doped indium oxide electrodes,
*
Corresponding authors. Tel./fax: +81 96 342 3656. E-mail address: [email protected] (M. Tominaga).
0022-0728/$ - see front matter Ó 2007 Published by Elsevier B.V. doi:10.1016/j.jelechem.2007.06.014
and cytochrome c3 on a mercury electrodes, were reported. In addition, various functionalized electrodes were developed for protein electrochemistry. In particular, self-assembled monolayer-modified electrodes have been widely developed for protein electrochemistry [4], and have been well-characterized by various surface analysis techniques [5–10]. Lipid film-modified electrodes also provide suitable conditions for direct electron transfer between proteins and electrodes [11,12]. Recently, direct heterogeneous electron transfer reactions between enzymes and electrodes have also been reported [2–29]. The direct electron transfer reactions on electrodes were achieved for heme-containing enzymes (such as cytochrome oxidases [12], cellobiose dehydrogenase [13,14], theophylline oxidase [15], alcohol dehydrogenase [13,16] and sulfite oxidase [17], peroxidase [13,18]), for
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multicopper enzymes (such as laccase [13,19,20], ascorbate oxidase [13,19], bilirubin oxidase [19,21], ceruloplasmin [19] and Cu efflux oxidase [22]), glucose oxidase on highly-oriented pyrolytic graphite and on carbon nanotubes [23,24], and superoxide dismutase [13,25]. However, for enzymes with large molecular weights, direct heterogeneous electron transfer reactions are still limited [26–29]. In general, the electron transfer rate of the enzyme on the electrode surface is slow or, cannot be detected due to the fact that the redox center of the enzyme is deeply buried within the protein shell, since the electron transfer rate is exponentially dependent on the distance between the redox active centers, as predicted by the Marcus theory [30,31]. Therefore, in order to shorten the electron transfer distance between the active center of the enzyme and the electrode, the proper molecular orientation of the enzyme on the electrode surface is one of the most important key requirements. In the present study, the electrode reactions of D-fructose dehydrogenase (FDH) from Gluconobacter sp. on the electrode surface were investigated. The obtained results clearly indicated that the proper molecular orientation of FDH on the electrode surface is of importance for direct heterogeneous electron transfer reactions of the enzyme on the electrode surface. FDH is a membrane-bound enzyme with a molecular weight of ca. 140 kDa containing flavin and heme c as prosthetic groups, which catalyze the oxidation of D-fructose to 2-keto-D-fructose [32]. The direct electron transfer reactions of FDH on electrode surfaces were reported in previous studies [33,34]. However, the investigations on proper molecular orientation for direct electron transfer reactions on an electrode are still insufficiently. In this study, using electrochemical and atomic force microscopic measurements, we demonstrated that the FDH complex decomposed in neutral and alkaline solutions, and in the case when no electrode reaction was observed. Furthermore, we indicated that the flavin-containing subunit in FDH accepted electrons from D-fructose and transferred the electrons to the heme c containing subunit, and then the direct electron transfer reaction of FDH occurred at that heme c site. 2. Experimental 2.1. Fructose dehydrogenase and materials D-Fructose dehydrogenase (FDH, EC 1.1.99.11, from Gluconobacter sp.) was purchased from Toyobo Co., Japan, and was used without further purification. Water was purified with a Millipore Milli-Q water system. All other chemicals were of analytical grade and were used as received. A plate of basal-plane highly oriented pyrolytic graphite (basal-plane HOPG, Panasonic graphite, PGX 05, electrode area: 0.44 cm2) was from Matsushita Electric Co., Japan. Prior to the use, the surface of the basal-plane HOPG was peeled off by adhesive tape to expose a fresh
basal-plane. A tin-doped indium oxide electrode (electrode area: 0.25 cm2, from Kinoene Optics Corp., Japan) was cleaned by ultra-sonication in 1% aqueous New-Vista (AIC Corp., Japan) solution, and an electrode with a fully hydrophilic surface (72 dyn cm 1 at 25 °C) was used as the working electrode. A plastic formed carbon plate (PFC, Mitsubishi Pencil Co., Japan, electrode area: 0.44 cm2) was used as the other working electrode. Prior to use, the PFC plate was polished with No. 400- and 2000-grid SiC papers, followed by sonication in Milli-Q water for 10 min. The FDH adsorbed electrode was prepared by immersion of the electrodes into a 100 mmol dm 3 phosphate solution (pH 5) of 1 unit ll 1 FDH for 1 min. Phosphate solutions with various pH values were prepared by a mixture of 100 mmol dm 3 NaH2PO4 and 100 mmol dm 3 Na2HPO4 solutions. 2.2. Instrumentation Cyclic voltammetric and differential pulse voltammetric measurements were performed with an Electrochemical Analyzer, Model 700B, CH-Instruments (Austin, TX, USA). An Ag/AgCl (saturated KCl) electrode and a Pt plate were used as the reference and auxiliary electrodes, respectively. All potentials are reported with respect to the Ag/AgCl (saturated KCl) electrode. Prior to the cyclic voltammetric measurements, the buffer solution was deaerated with high purity argon gas, and a positive pressure of argon gas was kept over the solution during all electrochemical experiments. X-ray photoelectron spectroscopy (XPS) measurements were performed using a Sigma Probe HA6000II, Thermo VG Scientific. This instrument uses a focused monochromatic Al Ka X-ray (1486.68 eV) source for excitation, a spherical section analyzer, and a 6-element multichannel detection system. A multimode NanoScope III (Digital Instruments) was utilized for tapping-mode atomic force microscopy (AFM) imaging. Standard phosphorus doped silicon cantilevers were used. The calibration was performed using an imaging standard grating sample. Tapping-mode AFM measurements were carried out under an air atmosphere. Fluorescent and UV–vis spectra were obtained using a Spectrofluorometer FP-6500, JASCO, Japan, and an UV–vis-NIR spectrophotometer, UV-3100, Shimadzu, Japan, respectively. Fluorescent images were obtained using a Variable mode imager, Typhoon 9400, Amersham Biosciences. The electrophoretic analyses were performed using polyacrylamide gel electrophoresis in Tris–glycine buffer in the presence of sodium dodecyl sulfate (SDS). A protein solution (100 mmol dm 3 Tris–HCl, pH 8.8) was made in 2% SDS and in 100 mmol dm 3 dithiothreitol, heated in a boiling water bath for 5 min, and then subjected to electrophoresis (20 mA per gel for ca. 80 min). The protein was stained with Coomassie brilliant blue.
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3. Results and discussion 3.1. Properties of D-fructose dehydrogenase It has been reported that D-fructose dehydrogenase solubilized and purified from the membrane fraction of glycerol-grown Gluconobacter industrius has a molecular weight of 140 kDa, with three components consisting of one flavin and one heme c as prosthetic groups [32]. To identify the FDH used in this study, spectroscopic measurements and electrophoretic analyses were carried out. Fig. 1a shows the UV–vis spectrum of the reduced form of FDH upon the addition of sodium dithionite as a reductant. Absorption peaks at 416, 522 and 553 nm were observed for the reduced form, which was a cytochrome c-like absorption spectrum. The obtained results, together with previous reports, indicated that the FDH complex contained heme c. Fig. 1b shows the fluorescent spectrum of FDH upon irradiation at 375 nm, which showed one peak at around 524 nm. This result clearly indicates that the FDH complex also contains flavin. The results of the electrophoretic analyses indicated that FDH was composed of three subunits with apparent molecular weights of 60.5 (±0.5), 48.3 (±0.3) and 18.7 (±0.2) kDa as shown in Fig. 2a. The sum of the molecular weight of the three subunits gave a total molecular weight of approximately 127.5 kDa, which was comparable to the previously reported value [32]. Further investigation using fluorescent scanning images from the SDS–gel electrophoresis of FDH indicated that the subunit with a molecular weight of 60.5 (±0.5) kDa contained flavin, whereas the subunit with a molecular weight of 48.3 (±0.3) kDa contained heme c. These results were in good agreement with previously reported results [32]. The function of the smallest subunit (molecular weight of ca. 18.7 kDa) remains unclear [32]. From the obtained results, the expected structure of FDH is shown in Fig. 2b.
Fig. 2. (a) SDS–gel electrophoreses of FDH. The molecular markers used were 14.3, 20.1, 29.0, 45.0, 66.4 and 97.2 kDa. (b) Schematic illustration for the expected structure of FDH.
3.2. Voltammetric behaviors of FDH adsorbed on electrodes Fig. 3 shows typical voltammetric curves obtained at the FDH adsorbed basal-plane HOPG, PFC and tin-doped indium oxide electrodes in phosphate solution (pH 5.0) in the presence and absence of 100 mmol dm 3 fructose. The catalytic current was observed from around 0.1 V at these electrodes, when fructose was present in the solution. No catalytic current was observed at the FDHunmodified electrodes. These observations indicated that direct heterogeneous electron transfer reactions of the adsorbed FDH occurred on the electrodes. The aforementioned behavior was similar to results reported by a previous study using FDH-adsorbed Ketjen black-modified glassy carbon electrodes [34]. The obtained catalytic currents for the adsorbed FDH did not increase, when the
Fig. 1. UV–vis (a) and fluorescent (b) spectra of FDH in a phosphate solution (pH 5). The excitation of the fluorescent spectra was 375 nm. The light pass lengths were 5 and 3 mm for the UV–vis and fluorescent spectra, respectively.
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Fig. 3. Typical cyclic voltammograms for FDH adsorbed onto basal-plane HOPG (0.44 cm 2) (a), PFC (0.44 cm 2) and tin-doped indium oxide (0.25 cm 2) electrodes in a phosphate solution (pH 5) in the presence (solid lines) and absence (broken lines) of 100 mmol dm 3 fructose. Potential sweep rate: 5 mV s 1.
additional fructose was present, indicating that the concentration of fructose was the substrate-saturated condition. To characterize the catalytic currents of the FDH adsorbed onto the electrode, the pH dependence was investigated. Fig. 4 shows the catalytic currents of the FDH adsorbed onto basal-plane HOPG electrode in various pH phosphate solutions. The catalytic current was observed in acidic solutions (pH 6 6). On the other hand, in neutral and alkaline (pH P 7) solutions, the catalytic current was not observed. These results indicated that neutral and alkaline (pH P 7) solutions were not suitable for the preparation of FDH adsorbed electrodes. The reason why the catalytic current was not observed in the case of the neutral and alkaline (pH P 7) solutions is unknown. However, it is possible that the FDH molecules could not adsorb onto the basal-plane HOPG electrode surface and/or the adsorbed FDH molecules desorbed into the solution from the electrode surface. To clarify the aforementioned possibility, XPS measurements were performed. Two samples were prepared as follows: the basal-plane HOPG electrodes were immersed into pH 5 or 8 phosphate solution containing 1 unit ll 1 FDH for 1 min, and were then rinsed with the same pH phos-
Fig. 4. pH dependence on the catalytic currents obtained from FDH adsorbed onto a basal-plane HOPG electrode in a phosphate solution (pH 5) in the presence of 100 mmol dm 3 fructose.
phate solution without FDH. The XPS results for both samples were very similar to each other. Peaks around 402 and 531 eV corresponding to N(1s) and O(1s), respectively, were observed [35], which were not observed before the FDH modification. These results clearly indicated that FDH was present on the electrode surface, even when the electrode was immersed into the pH 8 solution of FDH. The reason why the catalytic current was not obtained when FDH was modified onto electrodes in neutral and alkaline (pH P 7) solutions will be discussed in the following sections. 3.3. Atomic force microscopic measurements of FDH adsorbed on basal-plane HOPG plate Figs. 5 and 6 show AFM images of FDH molecules adsorbed onto basal-plane HOPG plate surfaces, when the plates were dipped in the phosphate solutions (pH 5 and 8) containing 0.15 unit ll 1 FDH for 5 s, followed by rinsing in the same pH solutions. Molecules with a height of ca. 7 nm were observed when the electrode was modified with FDH in the pH 5 solution. On the other hand, in the case of electrode modification in the pH 8 solution, molecules with a height of around 3 nm were observed. Taking into account that the FDH used was composed of three subunits with apparent molecular weights of 60.5 (±0.5) kDa containing flavin, 48.3 (±0.3) kDa containing heme c and 18.7 (±0.2) kDa as described above, the AFM results could be interpreted as follows. The molecules with around a 3 nm height obtained by electrode immersion into the pH 8 solution would be separated subunits produced by the decomposition of the FDH complex, as shown in Fig. 6d. In other words, the FDH complex was decomposed in the pH 8 solution, and then the separated subunits were adsorbed onto the electrode surface. We confirmed this decomposition behavior of FDH. When the electrode was immersed into a pH 5 solution of FDH followed by immersion into the pH 8 solution,
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Fig. 5. Tapping-mode AFM images for FDH adsorbed onto a basal-plane HOPG surface (a,c) when the plate was dipped in a phosphate solution (pH 5) containing 0.15 unit ll 1 FDH for 5 s. The cross-sectional view (b) corresponds to the line drawn in (a). The expected models for FDH molecules adsorbed onto the surface are shown in (d).
Fig. 6. Tapping-mode AFM images for FDH adsorbed onto a basal-plane HOPG surface (a,c), when the plate was dipped in a phosphate solution (pH 8) containing 0.15 unit ll 1 FDH for 5 s. The cross-sectional view (b) corresponds to the line drawn in (a). The expected models for the decomposed FDH subunits adsorbed onto the surface are shown in (d).
only molecules with around a 3 nm height were observed on the AFM images. This result clearly indicates that the FDH complex decomposes in neutral and alkaline
(pH P 7) solutions. On the other hand, the molecules with ca. 7 nm height obtained by electrode immersion into the pH 5 solution would be the FDH complex without
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decomposition, as shown in Fig. 5d. These AFM results explain the reason why the catalytic current could not be observed when FDH was modified onto the electrode in neutral and alkaline (pH P 7) solutions. 3.4. FDH complex orientation on the electrode surface for direct electron transfer reactions Fig. 7 shows differential pulse voltammograms for FDH adsorbed basal-plane HOPG electrode in various pH phosphate solutions. Two main peaks at ca. 0.15 V and in the potential range of 0.2 to 0.4 mV could be recognized. The peak at ca. 0.15 V was observed only in the acidic solution (pH 6 6). This peak would be concluded to represent an oxidation peak from the heme c-containing subunit in FDH, because the peak potential did not depend on the pH, and many proteins containing heme groups give a redox peak at around this potential [1–4]. The peak observed in the potential region of 0.2 to 0.4 V, was observed over the pH range from 4.4 to 8.0. This peak would be concluded to be an oxidation peak from the flavin-containing subunit in FDH, because the peak potential depended on the pH and flavin groups give a redox peak at around this potential [23,24]. In fact, the slope of a pH vs. oxidation peak potential plot was estimated to be ca. 59 mV pH 1 as shown in Fig. 7, which was in good agreement with the expected values for a 2-proton and 2-electron redox reaction of flavin [23,24]. The magnitude of the voltammetric peak for a heme was significantly small com-
pared to that for flavin, even though the number of electrons involved in the electrode reaction was half for a heme of that for flavin. This behavior may suggest the possibility of the preferential denaturing of heme site or the distribution of FDH orientation on the electrode surface. Fig. 8 shows the differential pulse voltammograms for FDH adsorbed onto basal-plane HOPG electrodes in the potential region from 0.2 to 0.25 V in a pH 5 phosphate solution in the presence and absence of 0.1 mol dm 3 fructose. The oxidation peak representing the heme c from FDH was observed at ca. 0.15 V when fructose was not present. When fructose was present, the catalytic oxidation current of fructose by FDH was obtained at an oxidation peak potential of 0.15 V for heme c. This result clearly indicated that the direct heterogeneous electron transfer reaction of FDH occurred at the heme c-containing subunit. In conclusion, the proposed electron relay model for
Fig. 8. Differential pulse voltammograms for FDH adsorbed basal-plane HOPG (a,b) and basal-plane HOPG (c) electrodes (0.44 cm2) in phosphate solution (pH 5) in the presence (a) and absence (b) of 100 mmol dm 3 fructose. Pulse amplitude: 50 mV; Sample width: 17 ms; pulse width: 50 ms; potential sweep rate: 5 mV s 1.
Fig. 7. Differential pulse voltammograms for FDH adsorbed basal-plane HOPG electrodes (0.44 cm2) in various pH phosphate solutions in the absence of fructose. Pulse amplitude: 50 mV; Sample width: 17 ms; pulse width: 50 ms; potential sweep rate: 5 mV s 1.
Fig. 9. Schematic illustration for the proposed electron relay model for the adsorbed FDH complex on the electrode surface.
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the FDH complex adsorbed onto the electrode surface is shown in Fig. 9. 4. Conclusions The direct heterogeneous electron transfer reactions and molecular orientation of D-fructose dehydrogenase adsorbed onto a basal-plane pyrolytic graphite electrode were investigated. From the results of the UV–vis, fluorescent spectroscopic measurements and electrophoretic analyses, it was clear that the FDH used in this study was composed of three subunits with apparent molecular weights of 60.5 (±0.5), 48.3 (±0.3) and 18.7 (±0.2) kDa. The subunits of molecular weights 60.5 (±0.5) and 48.3 (±0.3) kDa contained flavin and heme c, respectively. Catalytic oxidation currents based on the direct electron transfer of FDH adsorbed onto the basal-plane HOPG, PFC and tin-doped indium oxide electrodes were observed from a potential of around 0.1 V in phosphate solution (pH 5) in the presence of fructose as a substrate of FDH. The catalytic current for the FDH adsorbed onto the basal-plane HOPG electrode indicated a pH dependence. The catalytic oxidation currents were only observed in acidic solutions (pH 6 6), and were not observed in neutral and alkaline solutions. The reason why the catalytic current was not obtained in neutral and alkaline solutions was because the FDH complex decomposed in neutral and alkaline solutions, based on the results of AFM measurements. Differential pulse voltammetric investigations for FDH adsorbed onto the basal-plane HOPG electrodes, together with the AFM and electrophoresis results, indicated that the flavin-containing subunit in FDH accepts electrons from D-fructose and transfers these electrons to the heme c- containing subunit, and then a direct electron transfer reaction of FDH occurred at the heme c site. Finally, we suggest a possible adsorbed FDH complex orientation on the electrode surface for the direct electron transfer as shown in Fig. 9. The obtained results clearly indicated that the proper orientation of FDH on the electrode surface is of critical importance for successful direct electron transfer reactions of the enzyme on the electrode surface. Acknowledgements This work was supported in part by a Grand-in-Aid for Scientific Research (M.T.) from the Ministry of Education, Culture, Science, Sports and Technology of Japan. M.T. also acknowledges the Hosokawa Powder Technology Foundation. References [1] M.J. Eddowes, H.A.O. Hill, J. Chem. Soc., Chem. Commun. (1977) 771. [2] P. Yeh, T. Kuwana, Chem. Lett. (1977) 1145.
7
[3] K. Niki, T. Yagi, H. Inokuchi, K. Kimura, J. Am. Chem. Soc. 101 (1979) 3335. [4] I. Taniguchi, K. Toyosawa, H. Yamaguchi, K. Yasukouchi, J. Chem. Soc., Chem. Commun. (1982) 1032; I. Taniguchi, in: G. Dryhust (Ed.), Interfacial Electrochemistry of Promoter Modified Electrodes for the Rapid Electron Transfer of Cytochrome c, in Redox Chemistry and Interfacial Behavior of Biological Molecules, Plenum Press, New York, 1988, p. 113; S. Yoshimoto, Bull. Chem. Soc. Jpn. 8 (2006) 1167, and references cited therein. [5] W.C. Bigelow, D.L. Pickett, W.A. Zisman, J. Colloid Interf. Sci. 1 (1946) 513. [6] R.G. Nuzzo, D.L. Allara, J. Am. Chem. Soc. 105 (1983) 4481; D.L. Allara, R.G. Nuzzo, Langmuir 1 (1985) 45; D.L. Allara, R.G. Nuzzo, Langmuir 1 (1985) 52; L.H. Dubois, B.R. Zegarski, R.G. Nuzzo, J. Am. Chem. Soc. 112 (1990) 570; L.H. Dubois, B.R. Zegarski, R.G. Nuzzo, J. Chem. Phys. 98 (1993) 678. [7] N.E. Schlotter, M.D. Poter, T.B. Bright, D.L. Allara, Chem. Phys. Lett. 132 (1986) 93; E.B. Troughton, C.D. Bain, G.M. Whitesides, R.G. Nuzzo, D.L. Allara, M.D. Porter, Langmuir 4 (1988) 365. [8] M.D. Porter, T.B. Brigth, D.L. Allara, C.E.D. Chidsey, J. Am. Chem. Soc. 109 (1987) 3559; C.E.D. Chidsey, Science 251 (1991) 919; C.E.D. Chidsey, C.R. Bertozzi, T.M. Putvinski, A.M. Mujsce, J. Am. Chem. Soc. 112 (1990) 4301. [9] A. Ulman (Ed.), Characterization of Organic Thin Films, Butterworth-Heineman, Stoneham, MA, 1994; A. Ulman (Ed.), An Introduction to Ultrathin Organic Films from Langmuir–Blodgett to Self-Assembly, Academic, San Diego, CA, 1991; H.O. Finklea (Ed.), Electrochemistry of Organized Monolayers of Thiols and Related Molecules on Electrodes, Electroanalytical Chemistry: A Series of Advances, Electroanalytical Chemistry, vol. 19, Marcel Dekker, NY, 1996; A. Ulman, Chem. Rev. 96 (1996) 1533. [10] M.J. Tarlov, E.F. Bowden, J. Am. Chem. Soc. 113 (1991) 1847; Z.Q. Feng, S. Imabayashi, T. Kakiuchi, K. Niki, J. Chem. Soc., Faraday Trans. 93 (1997) 1367; L.J. Kepley, R.M. Crooks, A.J. Ricco, Anal. Chem. 64 (1992) 3191; C.E. Jordan, B.L. Frey, S. Korngruth, R.M. Corn, Langmuir 10 (1994) 3642; S. Terrettaz, J. Cheng, C.J. Miller, R.D. Guiles, J. Am. Chem. Soc. 118 (1996) 7857; G.B. Sigal, M. Mrksich, G.M. Whitesides, J. Am. Chem. Soc. 120 (1998) 3464; X.-Y. Hu, Y. Xiao, H.-Y. Chen, J. Electroanal. Chem. 466 (1999) 26; M. Tominaga, A. Ohira, A. Kubo, I. Taniguchi, M. Kunitake, Chem. Commun. (2004) 1518; M. Tominaga, S. Maetsu, A. Kubo, I. Taniguchi, J. Electroanal. Chem. 603 (2007) 203. [11] J.F. Rusling, Acc. Chem. Res. 31 (1998) 363, and references cited therein; P. Bianco, J. Haladjian, Electrochim. Acta 42 (1997) 587; T. Ferri, A. Poscia, R. Santucci, Bioelectrochem. Bioener. 44 (1998) 177; M. Tominaga, S. Hashimoto, A. Misaka, N. Nakashima, Anal. Chem. 71 (1999) 2790; M. Tominaga, S. Hashimoto, N. Nakashima, J. Electroanal. Chem. 561 (2004) 13; M. Tominaga, Electrochem. Commun. 4 (2002) 968; M. Tominaga, Electrochemistry 69 (2001) 937. [12] J.K. Cullison, F.M. Hawkridge, N. Nakashima, S. Yoshikawa, Langmuir 10 (1994) 877; J.D. Burgess, M.C. Rhoten, F.M. Hawkridge, J. Am. Chem. Soc. 120 (1998) 4488;
8
[13]
[14]
[15] [16]
[17] [18]
[19]
[20]
[21]
M. Tominaga et al. / Journal of Electroanalytical Chemistry 610 (2007) 1–8 J.D. Burgess, V.W. Jones, M.D. Porter, M.C. Rhoten, F.M. Hawkridge, Langmuir 14 (1998) 6628. L. Gorton, A. Lindgren, T. Larsson, F.D. Munteanu, T. Ruzgas, I. Gazaryan, Anal. Chim. Acta 400 (1999) 91, and references cited therein. A. Lindgren, L. Gorton, T. Ruzgas, U. Baminger, D. Haltrich, M. Schu¨lein, J. Electroanal. Chem. 496 (2001) 76; L. Stoica, N. Dimcheva, D. Haltrich, T. Ruzgas, L. Gorton, Biosens. Bioelectron. 20 (2005) 2010; L. Stoica, T. Ruzgas, R. Ludwig, D. Haltrich, L. Gorton, Langmuir 22 (2006) 10801; W. Harreither, V. Coman, R. Ludwig, D. Haltrich, L. Gorton, Electroanalysis 19 (2007) 172. A. Christenson, E. Dock, L. Gorton, T. Ruzgas, Biosens. Bioelectron. 20 (2004) 176. T. Ikeda, D. Kobayashi, S. Matsushita, T. Sagara, K. Niki, J. Electroanal. Chem. 362 (1993) 221; A. Ramanavicius, K. Habermuller, E. Csoregi, V. Laurinavicius, W. Schuhmann, Anal. Chem. 71 (1999) 3581; J. Razumiene, M. Niculescu, A. Ramanavicius, V. Laurinavicius, E. Cso¨regi, Electroanalysis 14 (2001) 43. E.E. Ferapontova, T. Ruzgas, L. Gorton, Anal. Chem. 75 (2003) 4841. E.E. Ferapontova, V.G. Grigorenko, A.M. Egorov, T. Bo¨rchers, T. Ruzgas, L. Gorton, J. Electroanal. Chem. 509 (2001) 19; S. Gaspar, H. Zimmermann, I. Gazaryan, E. Cso¨regi, W. Schuhmann, Electroanalysis 13 (2001) 284; R. Andreu, E.E. Ferapontova, L. Gorton, J.J. Calvente, J. Phys. Chem. B 111 (2007) 469. S. Shleev, J. Tkac, A. Christenson, T. Ruzgas, A.I. Yaropolov, J.W. Whittaker, L. Gorton, Biosens. Bioelectron. 20 (2005) 2517, and references cited therein. G. Gupta, V. Rajendran, P. Atanassov, Electroanalysis 16 (2004) 1182; S. Shleev, A. Christenson, V. Serezhenkov, D. Burbaev, A.I. Yaropolov, L. Gorton, T. Ruzgas, Biochem. J. 385 (2005) 745; M. Pita, S. Shleev, T. Ruzgas, V.M. Ferna´ndez, A.I. Yaropolov, L. Gorton, Electrochem. Commun. 8 (2006) 747; S. Shleev, M. Pita, A.I. Yaropolov, T. Ruzgas, L. Gorton, Electroanalysis 18 (2006) 1901. S. Tsujimura, T. Nakagawa, K. Kano, T. Ikeda, Electrochemistry 72 (2004) 437; S. Tsujimura, K. Kano, T. Ikeda, J. Electroanal. Chem. 576 (2005) 113; S. Shleev, A.E. Kasmi, T. Ruzgas, L. Gorton, Electrochem. Commun. 6 (2004) 934;
[22] [23] [24]
[25] [26] [27] [28] [29] [30] [31] [32]
[33] [34] [35]
S. Shleev, A.J. Wilkolazka, A. Khalunina, O. Morozova, A. Yaropolov, T. Ruzgas, L. Gorton, Bioelectrochemistry 67 (2005) 115; W. Zheng, Q. Li, L. Su, Y. Yan, J. Zhang, L. Mao, Electroanalysis 18 (2006) 587; M. Tominaga, M. Otani, M. Kishikawa, I. Taniguchi, Chem. Lett. 35 (2006) 1174. Y. Miura, S. Tsujimura, Y. Kamitaka, S. Kurose, K. Kataoka, T. Sakurai, K. Kano, Chem. Lett. 36 (2007) 132. G. Wang, N.M. Thai, S.-T. Yau, Electrochem. Commun. 8 (2006) 987. F. Patolsky, Y. Weizmann, I. Willner, Angew. Chem., Int. Ed. 43 (2004) 2113; K. Gong, Y. Yan, M. Zhang, L. Su, S. Xiong, L. Mao, Anal. Sci. 21 (2005) 1383, and references cited therein; J. Wang, Electroanalysis 17 (2005) 7, and references cited therein; J.J. Gooding, Electrochim. Acta 50 (2005) 3049, and references cited therein; N. Jia, L. Liu, Q. Zhou, L. Wang, M. Yan, Z. Jiang, Electrochim. Acta 51 (2005) 611; D. Ivnitski, B. Branch, P. Atanassov, C. Apblett, Electrochem. Commun. 8 (2006) 1204; C.E. Banks, A. Crossley, C. Salter, S.J. Wilkins, R.G. Compton, Angew. Chem., Int. Ed. 45 (2006) 2533. Y. Tian, L. Mao, T. Okajima, T. Ohsaka, Anal. Chem. 76 (2004) 4162. F.A. Armstrong, G.S. Wilson, Electrochim. Acta 45 (2000) 2623, and references cited therein. S.C. Barton, J. Gallaway, P. Atanassov, Chem. Rev. 104 (2004) 4867, and references cited therein. W. Zhang, G. Li, Anal. Sci. 20 (2004) 603. F.A. Armstrong, H.A.O. Hill, Acc. Chem. Res. 21 (1998) 407. R.A. Marcus, N. Sutin, Biochim. Biophys. Acta 811 (1985) 265. R.A. Marcus, Angew. Chem., Int. Ed. 32 (1993) 1111. M. Ameyama, E. Shinagawa, K. Matsushita, O. Adachi, J. Bacteriol. 145 (1981) 814; M. Ameyama, O. Adachi, Methods Enzymol. 89 (1982) 154. E.E. Ferapontova, L. Gorton, Bioelectrochemistry 66 (2005) 55. Y. Kamitaka, S. Tsujimura, K. Kano, Chem. Lett. 36 (2007) 218. J.F. Moulder, W.F. Stickle, P.E. Sobol, K.D. Bomben, in: J. Chastain, R.C. King Jr. (Eds.), A Reference Book of Standard Spectra for Identification and Interpretation of XPS Data, Handbook of X-ray Photoelectron Spectroscopy, Physical Electronics, Inc., Minnesota, 1995.
Electroanalytical Chemistry Journal of Electroanalytical Chemistry 610 (2007) 1–8 www.elsevier.com/locate/jelechem
Direct heterogeneous electron transfer reactions and molecular orientation of fructose dehydrogenase adsorbed onto pyrolytic graphite electrodes Masato Tominaga *, Chiharu Shirakihara, Isao Taniguchi
*
Graduate School of Science and Technology, Kumamoto University, Kumamoto 860-8555, Japan Received 25 April 2007; received in revised form 7 June 2007; accepted 19 June 2007 Available online 28 June 2007
Abstract Direct heterogeneous electron transfer reactions and the molecular orientation of D-fructose dehydrogenase (FDH, from Gluconobacter sp.) adsorbed onto electrodes were investigated. Catalytic oxidation currents based on the direct electron transfer reactions of FDH adsorbed onto basal-plane, highly oriented pyrolytic graphite (basal-plane HOPG), plastic formed carbon plate and tin-doped indium oxide electrodes were observed from a potential around 0.1 V (vs. Ag/AgCl/saturated KCl) in phosphate solution (pH 5.0) in the presence of fructose as a substrate for FDH. The catalytic current for the FDH adsorbed onto basal-plane HOPG electrodes indicated a pH dependence. The catalytic oxidation currents were only observed in acidic solutions (pH 6 6), and were not observed in neutral and alkaline solutions. The reason why the catalytic current was not obtained in neutral and alkaline solutions was because the FDH complex decomposed in neutral and alkaline solutions based on the AFM measurements. The differential pulse voltammetric investigations for FDH adsorbed onto basal-plane HOPG electrode together with the AFM and electrophoresis results indicated that the flavin-containing subunit in FDH accepts electrons from D-fructose, and transfers these electrons to the heme c-containing subunit, and then the direct electron transfer reaction of FDH occurred at the heme c site. Ó 2007 Published by Elsevier B.V. Keywords: Fructose dehydrogenase; Electron transfer reaction; Orientation; Catalytic current; Flavin; Heme
1. Introduction Direct heterogeneous electron transfer reactions between proteins and electrodes have been extensively studied from the viewpoints of both understanding the fundamental features of proteins, and for applications as biodevices, biosensors and enzyme-biofuel cell. The first direct electron transfer reactions of proteins were performed by the Hill and Eddowes [1], Kuwana and Yeh [2] and Niki et al. [3] groups, in 1977, 1977 and 1979, respectively, in which the reversible cyclic voltammetric responses of cytochrome c on gold and tin-doped indium oxide electrodes,
*
Corresponding authors. Tel./fax: +81 96 342 3656. E-mail address: [email protected] (M. Tominaga).
0022-0728/$ - see front matter Ó 2007 Published by Elsevier B.V. doi:10.1016/j.jelechem.2007.06.014
and cytochrome c3 on a mercury electrodes, were reported. In addition, various functionalized electrodes were developed for protein electrochemistry. In particular, self-assembled monolayer-modified electrodes have been widely developed for protein electrochemistry [4], and have been well-characterized by various surface analysis techniques [5–10]. Lipid film-modified electrodes also provide suitable conditions for direct electron transfer between proteins and electrodes [11,12]. Recently, direct heterogeneous electron transfer reactions between enzymes and electrodes have also been reported [2–29]. The direct electron transfer reactions on electrodes were achieved for heme-containing enzymes (such as cytochrome oxidases [12], cellobiose dehydrogenase [13,14], theophylline oxidase [15], alcohol dehydrogenase [13,16] and sulfite oxidase [17], peroxidase [13,18]), for
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multicopper enzymes (such as laccase [13,19,20], ascorbate oxidase [13,19], bilirubin oxidase [19,21], ceruloplasmin [19] and Cu efflux oxidase [22]), glucose oxidase on highly-oriented pyrolytic graphite and on carbon nanotubes [23,24], and superoxide dismutase [13,25]. However, for enzymes with large molecular weights, direct heterogeneous electron transfer reactions are still limited [26–29]. In general, the electron transfer rate of the enzyme on the electrode surface is slow or, cannot be detected due to the fact that the redox center of the enzyme is deeply buried within the protein shell, since the electron transfer rate is exponentially dependent on the distance between the redox active centers, as predicted by the Marcus theory [30,31]. Therefore, in order to shorten the electron transfer distance between the active center of the enzyme and the electrode, the proper molecular orientation of the enzyme on the electrode surface is one of the most important key requirements. In the present study, the electrode reactions of D-fructose dehydrogenase (FDH) from Gluconobacter sp. on the electrode surface were investigated. The obtained results clearly indicated that the proper molecular orientation of FDH on the electrode surface is of importance for direct heterogeneous electron transfer reactions of the enzyme on the electrode surface. FDH is a membrane-bound enzyme with a molecular weight of ca. 140 kDa containing flavin and heme c as prosthetic groups, which catalyze the oxidation of D-fructose to 2-keto-D-fructose [32]. The direct electron transfer reactions of FDH on electrode surfaces were reported in previous studies [33,34]. However, the investigations on proper molecular orientation for direct electron transfer reactions on an electrode are still insufficiently. In this study, using electrochemical and atomic force microscopic measurements, we demonstrated that the FDH complex decomposed in neutral and alkaline solutions, and in the case when no electrode reaction was observed. Furthermore, we indicated that the flavin-containing subunit in FDH accepted electrons from D-fructose and transferred the electrons to the heme c containing subunit, and then the direct electron transfer reaction of FDH occurred at that heme c site. 2. Experimental 2.1. Fructose dehydrogenase and materials D-Fructose dehydrogenase (FDH, EC 1.1.99.11, from Gluconobacter sp.) was purchased from Toyobo Co., Japan, and was used without further purification. Water was purified with a Millipore Milli-Q water system. All other chemicals were of analytical grade and were used as received. A plate of basal-plane highly oriented pyrolytic graphite (basal-plane HOPG, Panasonic graphite, PGX 05, electrode area: 0.44 cm2) was from Matsushita Electric Co., Japan. Prior to the use, the surface of the basal-plane HOPG was peeled off by adhesive tape to expose a fresh
basal-plane. A tin-doped indium oxide electrode (electrode area: 0.25 cm2, from Kinoene Optics Corp., Japan) was cleaned by ultra-sonication in 1% aqueous New-Vista (AIC Corp., Japan) solution, and an electrode with a fully hydrophilic surface (72 dyn cm 1 at 25 °C) was used as the working electrode. A plastic formed carbon plate (PFC, Mitsubishi Pencil Co., Japan, electrode area: 0.44 cm2) was used as the other working electrode. Prior to use, the PFC plate was polished with No. 400- and 2000-grid SiC papers, followed by sonication in Milli-Q water for 10 min. The FDH adsorbed electrode was prepared by immersion of the electrodes into a 100 mmol dm 3 phosphate solution (pH 5) of 1 unit ll 1 FDH for 1 min. Phosphate solutions with various pH values were prepared by a mixture of 100 mmol dm 3 NaH2PO4 and 100 mmol dm 3 Na2HPO4 solutions. 2.2. Instrumentation Cyclic voltammetric and differential pulse voltammetric measurements were performed with an Electrochemical Analyzer, Model 700B, CH-Instruments (Austin, TX, USA). An Ag/AgCl (saturated KCl) electrode and a Pt plate were used as the reference and auxiliary electrodes, respectively. All potentials are reported with respect to the Ag/AgCl (saturated KCl) electrode. Prior to the cyclic voltammetric measurements, the buffer solution was deaerated with high purity argon gas, and a positive pressure of argon gas was kept over the solution during all electrochemical experiments. X-ray photoelectron spectroscopy (XPS) measurements were performed using a Sigma Probe HA6000II, Thermo VG Scientific. This instrument uses a focused monochromatic Al Ka X-ray (1486.68 eV) source for excitation, a spherical section analyzer, and a 6-element multichannel detection system. A multimode NanoScope III (Digital Instruments) was utilized for tapping-mode atomic force microscopy (AFM) imaging. Standard phosphorus doped silicon cantilevers were used. The calibration was performed using an imaging standard grating sample. Tapping-mode AFM measurements were carried out under an air atmosphere. Fluorescent and UV–vis spectra were obtained using a Spectrofluorometer FP-6500, JASCO, Japan, and an UV–vis-NIR spectrophotometer, UV-3100, Shimadzu, Japan, respectively. Fluorescent images were obtained using a Variable mode imager, Typhoon 9400, Amersham Biosciences. The electrophoretic analyses were performed using polyacrylamide gel electrophoresis in Tris–glycine buffer in the presence of sodium dodecyl sulfate (SDS). A protein solution (100 mmol dm 3 Tris–HCl, pH 8.8) was made in 2% SDS and in 100 mmol dm 3 dithiothreitol, heated in a boiling water bath for 5 min, and then subjected to electrophoresis (20 mA per gel for ca. 80 min). The protein was stained with Coomassie brilliant blue.
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3. Results and discussion 3.1. Properties of D-fructose dehydrogenase It has been reported that D-fructose dehydrogenase solubilized and purified from the membrane fraction of glycerol-grown Gluconobacter industrius has a molecular weight of 140 kDa, with three components consisting of one flavin and one heme c as prosthetic groups [32]. To identify the FDH used in this study, spectroscopic measurements and electrophoretic analyses were carried out. Fig. 1a shows the UV–vis spectrum of the reduced form of FDH upon the addition of sodium dithionite as a reductant. Absorption peaks at 416, 522 and 553 nm were observed for the reduced form, which was a cytochrome c-like absorption spectrum. The obtained results, together with previous reports, indicated that the FDH complex contained heme c. Fig. 1b shows the fluorescent spectrum of FDH upon irradiation at 375 nm, which showed one peak at around 524 nm. This result clearly indicates that the FDH complex also contains flavin. The results of the electrophoretic analyses indicated that FDH was composed of three subunits with apparent molecular weights of 60.5 (±0.5), 48.3 (±0.3) and 18.7 (±0.2) kDa as shown in Fig. 2a. The sum of the molecular weight of the three subunits gave a total molecular weight of approximately 127.5 kDa, which was comparable to the previously reported value [32]. Further investigation using fluorescent scanning images from the SDS–gel electrophoresis of FDH indicated that the subunit with a molecular weight of 60.5 (±0.5) kDa contained flavin, whereas the subunit with a molecular weight of 48.3 (±0.3) kDa contained heme c. These results were in good agreement with previously reported results [32]. The function of the smallest subunit (molecular weight of ca. 18.7 kDa) remains unclear [32]. From the obtained results, the expected structure of FDH is shown in Fig. 2b.
Fig. 2. (a) SDS–gel electrophoreses of FDH. The molecular markers used were 14.3, 20.1, 29.0, 45.0, 66.4 and 97.2 kDa. (b) Schematic illustration for the expected structure of FDH.
3.2. Voltammetric behaviors of FDH adsorbed on electrodes Fig. 3 shows typical voltammetric curves obtained at the FDH adsorbed basal-plane HOPG, PFC and tin-doped indium oxide electrodes in phosphate solution (pH 5.0) in the presence and absence of 100 mmol dm 3 fructose. The catalytic current was observed from around 0.1 V at these electrodes, when fructose was present in the solution. No catalytic current was observed at the FDHunmodified electrodes. These observations indicated that direct heterogeneous electron transfer reactions of the adsorbed FDH occurred on the electrodes. The aforementioned behavior was similar to results reported by a previous study using FDH-adsorbed Ketjen black-modified glassy carbon electrodes [34]. The obtained catalytic currents for the adsorbed FDH did not increase, when the
Fig. 1. UV–vis (a) and fluorescent (b) spectra of FDH in a phosphate solution (pH 5). The excitation of the fluorescent spectra was 375 nm. The light pass lengths were 5 and 3 mm for the UV–vis and fluorescent spectra, respectively.
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Fig. 3. Typical cyclic voltammograms for FDH adsorbed onto basal-plane HOPG (0.44 cm 2) (a), PFC (0.44 cm 2) and tin-doped indium oxide (0.25 cm 2) electrodes in a phosphate solution (pH 5) in the presence (solid lines) and absence (broken lines) of 100 mmol dm 3 fructose. Potential sweep rate: 5 mV s 1.
additional fructose was present, indicating that the concentration of fructose was the substrate-saturated condition. To characterize the catalytic currents of the FDH adsorbed onto the electrode, the pH dependence was investigated. Fig. 4 shows the catalytic currents of the FDH adsorbed onto basal-plane HOPG electrode in various pH phosphate solutions. The catalytic current was observed in acidic solutions (pH 6 6). On the other hand, in neutral and alkaline (pH P 7) solutions, the catalytic current was not observed. These results indicated that neutral and alkaline (pH P 7) solutions were not suitable for the preparation of FDH adsorbed electrodes. The reason why the catalytic current was not observed in the case of the neutral and alkaline (pH P 7) solutions is unknown. However, it is possible that the FDH molecules could not adsorb onto the basal-plane HOPG electrode surface and/or the adsorbed FDH molecules desorbed into the solution from the electrode surface. To clarify the aforementioned possibility, XPS measurements were performed. Two samples were prepared as follows: the basal-plane HOPG electrodes were immersed into pH 5 or 8 phosphate solution containing 1 unit ll 1 FDH for 1 min, and were then rinsed with the same pH phos-
Fig. 4. pH dependence on the catalytic currents obtained from FDH adsorbed onto a basal-plane HOPG electrode in a phosphate solution (pH 5) in the presence of 100 mmol dm 3 fructose.
phate solution without FDH. The XPS results for both samples were very similar to each other. Peaks around 402 and 531 eV corresponding to N(1s) and O(1s), respectively, were observed [35], which were not observed before the FDH modification. These results clearly indicated that FDH was present on the electrode surface, even when the electrode was immersed into the pH 8 solution of FDH. The reason why the catalytic current was not obtained when FDH was modified onto electrodes in neutral and alkaline (pH P 7) solutions will be discussed in the following sections. 3.3. Atomic force microscopic measurements of FDH adsorbed on basal-plane HOPG plate Figs. 5 and 6 show AFM images of FDH molecules adsorbed onto basal-plane HOPG plate surfaces, when the plates were dipped in the phosphate solutions (pH 5 and 8) containing 0.15 unit ll 1 FDH for 5 s, followed by rinsing in the same pH solutions. Molecules with a height of ca. 7 nm were observed when the electrode was modified with FDH in the pH 5 solution. On the other hand, in the case of electrode modification in the pH 8 solution, molecules with a height of around 3 nm were observed. Taking into account that the FDH used was composed of three subunits with apparent molecular weights of 60.5 (±0.5) kDa containing flavin, 48.3 (±0.3) kDa containing heme c and 18.7 (±0.2) kDa as described above, the AFM results could be interpreted as follows. The molecules with around a 3 nm height obtained by electrode immersion into the pH 8 solution would be separated subunits produced by the decomposition of the FDH complex, as shown in Fig. 6d. In other words, the FDH complex was decomposed in the pH 8 solution, and then the separated subunits were adsorbed onto the electrode surface. We confirmed this decomposition behavior of FDH. When the electrode was immersed into a pH 5 solution of FDH followed by immersion into the pH 8 solution,
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Fig. 5. Tapping-mode AFM images for FDH adsorbed onto a basal-plane HOPG surface (a,c) when the plate was dipped in a phosphate solution (pH 5) containing 0.15 unit ll 1 FDH for 5 s. The cross-sectional view (b) corresponds to the line drawn in (a). The expected models for FDH molecules adsorbed onto the surface are shown in (d).
Fig. 6. Tapping-mode AFM images for FDH adsorbed onto a basal-plane HOPG surface (a,c), when the plate was dipped in a phosphate solution (pH 8) containing 0.15 unit ll 1 FDH for 5 s. The cross-sectional view (b) corresponds to the line drawn in (a). The expected models for the decomposed FDH subunits adsorbed onto the surface are shown in (d).
only molecules with around a 3 nm height were observed on the AFM images. This result clearly indicates that the FDH complex decomposes in neutral and alkaline
(pH P 7) solutions. On the other hand, the molecules with ca. 7 nm height obtained by electrode immersion into the pH 5 solution would be the FDH complex without
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decomposition, as shown in Fig. 5d. These AFM results explain the reason why the catalytic current could not be observed when FDH was modified onto the electrode in neutral and alkaline (pH P 7) solutions. 3.4. FDH complex orientation on the electrode surface for direct electron transfer reactions Fig. 7 shows differential pulse voltammograms for FDH adsorbed basal-plane HOPG electrode in various pH phosphate solutions. Two main peaks at ca. 0.15 V and in the potential range of 0.2 to 0.4 mV could be recognized. The peak at ca. 0.15 V was observed only in the acidic solution (pH 6 6). This peak would be concluded to represent an oxidation peak from the heme c-containing subunit in FDH, because the peak potential did not depend on the pH, and many proteins containing heme groups give a redox peak at around this potential [1–4]. The peak observed in the potential region of 0.2 to 0.4 V, was observed over the pH range from 4.4 to 8.0. This peak would be concluded to be an oxidation peak from the flavin-containing subunit in FDH, because the peak potential depended on the pH and flavin groups give a redox peak at around this potential [23,24]. In fact, the slope of a pH vs. oxidation peak potential plot was estimated to be ca. 59 mV pH 1 as shown in Fig. 7, which was in good agreement with the expected values for a 2-proton and 2-electron redox reaction of flavin [23,24]. The magnitude of the voltammetric peak for a heme was significantly small com-
pared to that for flavin, even though the number of electrons involved in the electrode reaction was half for a heme of that for flavin. This behavior may suggest the possibility of the preferential denaturing of heme site or the distribution of FDH orientation on the electrode surface. Fig. 8 shows the differential pulse voltammograms for FDH adsorbed onto basal-plane HOPG electrodes in the potential region from 0.2 to 0.25 V in a pH 5 phosphate solution in the presence and absence of 0.1 mol dm 3 fructose. The oxidation peak representing the heme c from FDH was observed at ca. 0.15 V when fructose was not present. When fructose was present, the catalytic oxidation current of fructose by FDH was obtained at an oxidation peak potential of 0.15 V for heme c. This result clearly indicated that the direct heterogeneous electron transfer reaction of FDH occurred at the heme c-containing subunit. In conclusion, the proposed electron relay model for
Fig. 8. Differential pulse voltammograms for FDH adsorbed basal-plane HOPG (a,b) and basal-plane HOPG (c) electrodes (0.44 cm2) in phosphate solution (pH 5) in the presence (a) and absence (b) of 100 mmol dm 3 fructose. Pulse amplitude: 50 mV; Sample width: 17 ms; pulse width: 50 ms; potential sweep rate: 5 mV s 1.
Fig. 7. Differential pulse voltammograms for FDH adsorbed basal-plane HOPG electrodes (0.44 cm2) in various pH phosphate solutions in the absence of fructose. Pulse amplitude: 50 mV; Sample width: 17 ms; pulse width: 50 ms; potential sweep rate: 5 mV s 1.
Fig. 9. Schematic illustration for the proposed electron relay model for the adsorbed FDH complex on the electrode surface.
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the FDH complex adsorbed onto the electrode surface is shown in Fig. 9. 4. Conclusions The direct heterogeneous electron transfer reactions and molecular orientation of D-fructose dehydrogenase adsorbed onto a basal-plane pyrolytic graphite electrode were investigated. From the results of the UV–vis, fluorescent spectroscopic measurements and electrophoretic analyses, it was clear that the FDH used in this study was composed of three subunits with apparent molecular weights of 60.5 (±0.5), 48.3 (±0.3) and 18.7 (±0.2) kDa. The subunits of molecular weights 60.5 (±0.5) and 48.3 (±0.3) kDa contained flavin and heme c, respectively. Catalytic oxidation currents based on the direct electron transfer of FDH adsorbed onto the basal-plane HOPG, PFC and tin-doped indium oxide electrodes were observed from a potential of around 0.1 V in phosphate solution (pH 5) in the presence of fructose as a substrate of FDH. The catalytic current for the FDH adsorbed onto the basal-plane HOPG electrode indicated a pH dependence. The catalytic oxidation currents were only observed in acidic solutions (pH 6 6), and were not observed in neutral and alkaline solutions. The reason why the catalytic current was not obtained in neutral and alkaline solutions was because the FDH complex decomposed in neutral and alkaline solutions, based on the results of AFM measurements. Differential pulse voltammetric investigations for FDH adsorbed onto the basal-plane HOPG electrodes, together with the AFM and electrophoresis results, indicated that the flavin-containing subunit in FDH accepts electrons from D-fructose and transfers these electrons to the heme c- containing subunit, and then a direct electron transfer reaction of FDH occurred at the heme c site. Finally, we suggest a possible adsorbed FDH complex orientation on the electrode surface for the direct electron transfer as shown in Fig. 9. The obtained results clearly indicated that the proper orientation of FDH on the electrode surface is of critical importance for successful direct electron transfer reactions of the enzyme on the electrode surface. Acknowledgements This work was supported in part by a Grand-in-Aid for Scientific Research (M.T.) from the Ministry of Education, Culture, Science, Sports and Technology of Japan. M.T. also acknowledges the Hosokawa Powder Technology Foundation. References [1] M.J. Eddowes, H.A.O. Hill, J. Chem. Soc., Chem. Commun. (1977) 771. [2] P. Yeh, T. Kuwana, Chem. Lett. (1977) 1145.
7
[3] K. Niki, T. Yagi, H. Inokuchi, K. Kimura, J. Am. Chem. Soc. 101 (1979) 3335. [4] I. Taniguchi, K. Toyosawa, H. Yamaguchi, K. Yasukouchi, J. Chem. Soc., Chem. Commun. (1982) 1032; I. Taniguchi, in: G. Dryhust (Ed.), Interfacial Electrochemistry of Promoter Modified Electrodes for the Rapid Electron Transfer of Cytochrome c, in Redox Chemistry and Interfacial Behavior of Biological Molecules, Plenum Press, New York, 1988, p. 113; S. Yoshimoto, Bull. Chem. Soc. Jpn. 8 (2006) 1167, and references cited therein. [5] W.C. Bigelow, D.L. Pickett, W.A. Zisman, J. Colloid Interf. Sci. 1 (1946) 513. [6] R.G. Nuzzo, D.L. Allara, J. Am. Chem. Soc. 105 (1983) 4481; D.L. Allara, R.G. Nuzzo, Langmuir 1 (1985) 45; D.L. Allara, R.G. Nuzzo, Langmuir 1 (1985) 52; L.H. Dubois, B.R. Zegarski, R.G. Nuzzo, J. Am. Chem. Soc. 112 (1990) 570; L.H. Dubois, B.R. Zegarski, R.G. Nuzzo, J. Chem. Phys. 98 (1993) 678. [7] N.E. Schlotter, M.D. Poter, T.B. Bright, D.L. Allara, Chem. Phys. Lett. 132 (1986) 93; E.B. Troughton, C.D. Bain, G.M. Whitesides, R.G. Nuzzo, D.L. Allara, M.D. Porter, Langmuir 4 (1988) 365. [8] M.D. Porter, T.B. Brigth, D.L. Allara, C.E.D. Chidsey, J. Am. Chem. Soc. 109 (1987) 3559; C.E.D. Chidsey, Science 251 (1991) 919; C.E.D. Chidsey, C.R. Bertozzi, T.M. Putvinski, A.M. Mujsce, J. Am. Chem. Soc. 112 (1990) 4301. [9] A. Ulman (Ed.), Characterization of Organic Thin Films, Butterworth-Heineman, Stoneham, MA, 1994; A. Ulman (Ed.), An Introduction to Ultrathin Organic Films from Langmuir–Blodgett to Self-Assembly, Academic, San Diego, CA, 1991; H.O. Finklea (Ed.), Electrochemistry of Organized Monolayers of Thiols and Related Molecules on Electrodes, Electroanalytical Chemistry: A Series of Advances, Electroanalytical Chemistry, vol. 19, Marcel Dekker, NY, 1996; A. Ulman, Chem. Rev. 96 (1996) 1533. [10] M.J. Tarlov, E.F. Bowden, J. Am. Chem. Soc. 113 (1991) 1847; Z.Q. Feng, S. Imabayashi, T. Kakiuchi, K. Niki, J. Chem. Soc., Faraday Trans. 93 (1997) 1367; L.J. Kepley, R.M. Crooks, A.J. Ricco, Anal. Chem. 64 (1992) 3191; C.E. Jordan, B.L. Frey, S. Korngruth, R.M. Corn, Langmuir 10 (1994) 3642; S. Terrettaz, J. Cheng, C.J. Miller, R.D. Guiles, J. Am. Chem. Soc. 118 (1996) 7857; G.B. Sigal, M. Mrksich, G.M. Whitesides, J. Am. Chem. Soc. 120 (1998) 3464; X.-Y. Hu, Y. Xiao, H.-Y. Chen, J. Electroanal. Chem. 466 (1999) 26; M. Tominaga, A. Ohira, A. Kubo, I. Taniguchi, M. Kunitake, Chem. Commun. (2004) 1518; M. Tominaga, S. Maetsu, A. Kubo, I. Taniguchi, J. Electroanal. Chem. 603 (2007) 203. [11] J.F. Rusling, Acc. Chem. Res. 31 (1998) 363, and references cited therein; P. Bianco, J. Haladjian, Electrochim. Acta 42 (1997) 587; T. Ferri, A. Poscia, R. Santucci, Bioelectrochem. Bioener. 44 (1998) 177; M. Tominaga, S. Hashimoto, A. Misaka, N. Nakashima, Anal. Chem. 71 (1999) 2790; M. Tominaga, S. Hashimoto, N. Nakashima, J. Electroanal. Chem. 561 (2004) 13; M. Tominaga, Electrochem. Commun. 4 (2002) 968; M. Tominaga, Electrochemistry 69 (2001) 937. [12] J.K. Cullison, F.M. Hawkridge, N. Nakashima, S. Yoshikawa, Langmuir 10 (1994) 877; J.D. Burgess, M.C. Rhoten, F.M. Hawkridge, J. Am. Chem. Soc. 120 (1998) 4488;
8
[13]
[14]
[15] [16]
[17] [18]
[19]
[20]
[21]
M. Tominaga et al. / Journal of Electroanalytical Chemistry 610 (2007) 1–8 J.D. Burgess, V.W. Jones, M.D. Porter, M.C. Rhoten, F.M. Hawkridge, Langmuir 14 (1998) 6628. L. Gorton, A. Lindgren, T. Larsson, F.D. Munteanu, T. Ruzgas, I. Gazaryan, Anal. Chim. Acta 400 (1999) 91, and references cited therein. A. Lindgren, L. Gorton, T. Ruzgas, U. Baminger, D. Haltrich, M. Schu¨lein, J. Electroanal. Chem. 496 (2001) 76; L. Stoica, N. Dimcheva, D. Haltrich, T. Ruzgas, L. Gorton, Biosens. Bioelectron. 20 (2005) 2010; L. Stoica, T. Ruzgas, R. Ludwig, D. Haltrich, L. Gorton, Langmuir 22 (2006) 10801; W. Harreither, V. Coman, R. Ludwig, D. Haltrich, L. Gorton, Electroanalysis 19 (2007) 172. A. Christenson, E. Dock, L. Gorton, T. Ruzgas, Biosens. Bioelectron. 20 (2004) 176. T. Ikeda, D. Kobayashi, S. Matsushita, T. Sagara, K. Niki, J. Electroanal. Chem. 362 (1993) 221; A. Ramanavicius, K. Habermuller, E. Csoregi, V. Laurinavicius, W. Schuhmann, Anal. Chem. 71 (1999) 3581; J. Razumiene, M. Niculescu, A. Ramanavicius, V. Laurinavicius, E. Cso¨regi, Electroanalysis 14 (2001) 43. E.E. Ferapontova, T. Ruzgas, L. Gorton, Anal. Chem. 75 (2003) 4841. E.E. Ferapontova, V.G. Grigorenko, A.M. Egorov, T. Bo¨rchers, T. Ruzgas, L. Gorton, J. Electroanal. Chem. 509 (2001) 19; S. Gaspar, H. Zimmermann, I. Gazaryan, E. Cso¨regi, W. Schuhmann, Electroanalysis 13 (2001) 284; R. Andreu, E.E. Ferapontova, L. Gorton, J.J. Calvente, J. Phys. Chem. B 111 (2007) 469. S. Shleev, J. Tkac, A. Christenson, T. Ruzgas, A.I. Yaropolov, J.W. Whittaker, L. Gorton, Biosens. Bioelectron. 20 (2005) 2517, and references cited therein. G. Gupta, V. Rajendran, P. Atanassov, Electroanalysis 16 (2004) 1182; S. Shleev, A. Christenson, V. Serezhenkov, D. Burbaev, A.I. Yaropolov, L. Gorton, T. Ruzgas, Biochem. J. 385 (2005) 745; M. Pita, S. Shleev, T. Ruzgas, V.M. Ferna´ndez, A.I. Yaropolov, L. Gorton, Electrochem. Commun. 8 (2006) 747; S. Shleev, M. Pita, A.I. Yaropolov, T. Ruzgas, L. Gorton, Electroanalysis 18 (2006) 1901. S. Tsujimura, T. Nakagawa, K. Kano, T. Ikeda, Electrochemistry 72 (2004) 437; S. Tsujimura, K. Kano, T. Ikeda, J. Electroanal. Chem. 576 (2005) 113; S. Shleev, A.E. Kasmi, T. Ruzgas, L. Gorton, Electrochem. Commun. 6 (2004) 934;
[22] [23] [24]
[25] [26] [27] [28] [29] [30] [31] [32]
[33] [34] [35]
S. Shleev, A.J. Wilkolazka, A. Khalunina, O. Morozova, A. Yaropolov, T. Ruzgas, L. Gorton, Bioelectrochemistry 67 (2005) 115; W. Zheng, Q. Li, L. Su, Y. Yan, J. Zhang, L. Mao, Electroanalysis 18 (2006) 587; M. Tominaga, M. Otani, M. Kishikawa, I. Taniguchi, Chem. Lett. 35 (2006) 1174. Y. Miura, S. Tsujimura, Y. Kamitaka, S. Kurose, K. Kataoka, T. Sakurai, K. Kano, Chem. Lett. 36 (2007) 132. G. Wang, N.M. Thai, S.-T. Yau, Electrochem. Commun. 8 (2006) 987. F. Patolsky, Y. Weizmann, I. Willner, Angew. Chem., Int. Ed. 43 (2004) 2113; K. Gong, Y. Yan, M. Zhang, L. Su, S. Xiong, L. Mao, Anal. Sci. 21 (2005) 1383, and references cited therein; J. Wang, Electroanalysis 17 (2005) 7, and references cited therein; J.J. Gooding, Electrochim. Acta 50 (2005) 3049, and references cited therein; N. Jia, L. Liu, Q. Zhou, L. Wang, M. Yan, Z. Jiang, Electrochim. Acta 51 (2005) 611; D. Ivnitski, B. Branch, P. Atanassov, C. Apblett, Electrochem. Commun. 8 (2006) 1204; C.E. Banks, A. Crossley, C. Salter, S.J. Wilkins, R.G. Compton, Angew. Chem., Int. Ed. 45 (2006) 2533. Y. Tian, L. Mao, T. Okajima, T. Ohsaka, Anal. Chem. 76 (2004) 4162. F.A. Armstrong, G.S. Wilson, Electrochim. Acta 45 (2000) 2623, and references cited therein. S.C. Barton, J. Gallaway, P. Atanassov, Chem. Rev. 104 (2004) 4867, and references cited therein. W. Zhang, G. Li, Anal. Sci. 20 (2004) 603. F.A. Armstrong, H.A.O. Hill, Acc. Chem. Res. 21 (1998) 407. R.A. Marcus, N. Sutin, Biochim. Biophys. Acta 811 (1985) 265. R.A. Marcus, Angew. Chem., Int. Ed. 32 (1993) 1111. M. Ameyama, E. Shinagawa, K. Matsushita, O. Adachi, J. Bacteriol. 145 (1981) 814; M. Ameyama, O. Adachi, Methods Enzymol. 89 (1982) 154. E.E. Ferapontova, L. Gorton, Bioelectrochemistry 66 (2005) 55. Y. Kamitaka, S. Tsujimura, K. Kano, Chem. Lett. 36 (2007) 218. J.F. Moulder, W.F. Stickle, P.E. Sobol, K.D. Bomben, in: J. Chastain, R.C. King Jr. (Eds.), A Reference Book of Standard Spectra for Identification and Interpretation of XPS Data, Handbook of X-ray Photoelectron Spectroscopy, Physical Electronics, Inc., Minnesota, 1995.