Direct Heterogeneous Electron Transfer Reactions ofBacillus halodurans Bacterial Blue Multicopper Oxidase

Electrochemistry Communications 8 (2006) 747–753 www.elsevier.com/locate/elecom Direct heterogeneous electron transfer reactions of fungal laccases at bare and thiol-modified gold electrodes Marcos Pita a, Sergey Shleev b,c,d,*, Tautgirdas Ruzgas d, Vı́ctor M. Fernández a, Alexander I. Yaropolov c, Lo Gorton b a c Instituto de Catálisis y Petroleoquı́mica CSIC, Marie Curie 2, 28049 Madrid, Spain b Lund University, Department of Analytical Chemistry, SE-221 00 Lund, Sweden Institute of Biochemistry, Laboratory of Chemical Enzymology, 119071 Moscow, Russia d Malmö University, Faculty of Health and Society, SE-205 06 Malmö, Sweden Received 17 February 2006; received in revised form 2 March 2006; accepted 6 March 2006 Available online 4 April 2006 Abstract Mediatorless (direct) electron transfer between bare and thiol-modified gold electrodes and fungal laccases from different sources has been demonstrated. The electrochemical activity of the enzymes from basidiomycetes Trametes hirsuta, Trametes ochracea, and Cerrena maxima under aerobic and anaerobic conditions can clearly be observed using cyclic voltammetry and spectroelectrochemistry. Bioelectroreduction of oxygen by T. hirsuta laccase immobilized on amino-thiophenol-modified gold electrodes, starting at +625 mV vs. NHE, is demonstrated and differences in bioelectrocatalysis of the enzyme immobilized on bare and thiol-modified electrodes are shown. It was found that hydrogen peroxide was one of the products of oxygen electroreduction on gold electrodes modified with fungal laccases, whereas no significant peroxide formation was observed for T. hirsuta laccase immobilised on thiol-modified gold electrodes. Thus, a hypothesis about two different mechanisms of oxygen electroreduction by fungal laccases adsorbed on bare and thiol-modified electrodes is proposed.  2006 Elsevier B.V. All rights reserved. Keywords: Laccase; Redox potential; T1, T2, and T3 sites; Gold electrode; Cyclic voltammetry 1. Introduction Studies of direct electron transfer (DET) reactions between laccases and electrodes yield important fundamental information about the thermodynamic and kinetic aspects of direct oxygen reduction to water, with no formation of highly toxic reactive oxygen species [1,2]. Moreover, the understanding of the heterogeneous reactions facilitates practical applications of laccases, such as creation of biosensors [3–6] and biofuel cells [7–9]. Laccase (benzenediol: oxygen oxidoreductases, EC 1.10.3.2) catalyses the oxidation of different kind of phe- * Abbreviations: DET, direct electron transfer; ET, electron transfer. Corresponding author. Tel.: +46 46 222 8191; fax: +46 46 222 4544. E-mail address: [email protected] (S. Shleev). 1388-2481/$ - see front matter  2006 Elsevier B.V. All rights reserved. doi:10.1016/j.elecom.2006.03.008 nols, amines and lignins, as well as some inorganic ions, coupled to the reduction of molecular oxygen to water [10,11]. The enzymes contain four metal ions historically classified into three types, e.g., T1, T2, T3, according to their spectral characteristics [11]. The T1 copper is responsible of the enzyme blue colour with a characteristic lightabsorbance band with a wavelength around 610 nm. The T2 copper cannot be detected by UV–vis spectroscopy, however, it generates a characteristic EPR-signal [12,13]. The bi-nuclear T3 copper is diamagnetic, so it is not EPR detectable. However, the T3 copper displays a spectral absorbance shoulder in the region of 330 nm [11] and it has a characteristic fluorescence spectrum [14,15]. The most remarkable characteristic of laccase is the redox potentials of the T1 Cu sites, which have been determined by potentiometric titrations with redox mediators 748 M. Pita et al. / Electrochemistry Communications 8 (2006) 747–753 for a great number of different laccases. The values of the redox potentials of the T1 site vary between 430 and 790 mV vs. NHE [13,16–19]. Laccases are classified into three groups according to the redox potential of the T1 site and their primary structures, namely low, middle, and high redox potential enzymes [2,13]. The last group is of special interest in biotechnology, e.g., for different bleaching [20– 22] and bioremediation processes [23,24], as well as for biosensors [25–27] and biofuel cells [28–30]. Gold is often used as electrode material due to its stability as a noble metal, and its many possibilities for the immobilization of biomolecules. However, until present, few reports exist concerning electrochemical studies of high potential laccases at gold electrodes [31–35]. It has been shown that different fungal laccases exhibited electrochemical activity on bare and thiol-modified electrodes with a midpoint potential close to 400 mV vs. NHE, which is too far away from the redox potential of the T1 site of fungal enzymes [32,33,35]. Some authors [32] concluded that the T1 site is in DET with gold, despite a considerable difference between the formal potential measured with cyclic voltammetry (410 mV) and the redox potential of the T1 site (780 mV) determined with redox titration. In a previous work another explanation is suggested, namely that the T2 site of laccase is in DET contact with gold causing the determined low redox potential values of the enzymes [35] and the hypothesis that the redox potential of the T2 copper site in low- and high-potential laccases from different sources might be very similar, i.e. approximately 400 mV, has been proposed [2,35]. The recently published results related to bioelectrocatalysis of plant (Rhus vernicifera) and fungal (Trametes hirsuta) laccases immobilised on thiol-modified gold electrodes showed almost similar redox potentials of oxygen electroreduction for both enzymes, while the absence of any electrochemical activity for plant and fungal laccases was found under anaerobic conditions [34]. Thus, the data on the ET process between high redox potential laccases and gold are rather limited and some results are not fully understood. Therefore we wanted to investigate the possibility of achieving DET for several fungal laccases from different basidiomycetes under aerobic and anaerobic conditions using bare and thiol-modified gold electrodes, as a way to gain insight into the mechanism of oxygen electroreduction by fungal laccases. 2. Experimental 2.1. Chemicals Sulphuric acid, H2O2, Na2HPO4, KH2PO4, EDTA, NaF, and KCl were obtained from Merck (Darmstadt, Germany). Cystamine, L-cysteine, 4-aminothiophenol, and 2,2 0 -azinobis-(3-ethylbenzthiazoline-6-sulphonate) (ABTS) were from Sigma Chem. Comp. (St. Louis, MO, USA). Absolute ethanol (99.7%) was from Solveco Chemicals AB (Täby, Sweden). All chemicals were of analytical grade. The buffers were prepared using water (18.2 MX) purified with a Milli-Q system (Millipore, Milford, CT, USA). 2.2. Enzymes The following basidiomycetes Trametes hirsuta (Wulfen) Pilát, Trametes ochracea (Pers.) Gib. and Ryvarden, and Cerrena maxima (Mont.) A. David & Rajchenb were used for laccase production and these strains were obtained from the laboratory collection of the Moscow State University of Engineering Ecology (Moscow, Russia). Detailed accounts of the preparative scale production of these laccases and their biochemical characterisation have been previously published [13]. Homogeneous preparations of these laccases were stored in 0.1 M phosphate buffer, pH 6.5, at 18 C. Horseradish peroxidase (HRP, Type VI, 300 units/mg, MW 40.0 kDa, RZ 2.7) was purchased from Sigma. 2.3. Electrochemical measurements 2.3.1. Spectroelectrochemical studies of laccases Electrochemically driven mediatorless redox transformations of the enzymes were carried out using a spectroelectrochemical cell consisting on a gold capillary electrode with a diameter of 0.8 mm and total volume of 0.7 ll. The design of the cell was described elsewhere [36]. The potential of the spectroelectrochemical cell was controlled by a three-electrode potentiostat (BAS LC-3E, Bioanalytical Systems, West Lafayette, IN, USA). Cyclic voltammograms of laccase in the gold capillary electrode were recorded using a three-electrode potentiostat (BAS CV-100 W Electrochemical Analyser with BAS CV-100W software v. 2.1.). In both steady state and the CV measurements an Ag|AgCl|KClsat reference electrode (200 mV vs. NHE) and a platinum counter electrode were used. The absorbance spectra were monitored with PC2000-UV– VIS, a miniature fibre optic spectrometer from Ocean Optics (Dunedin, FL, USA) with an effective range between 200 and 1100 nm. The pre-treatment of the gold capillary working electrode of the spectroelectrochemical cell was carried out as described previously [35]. 2.3.2. Electrochemical measurements on planar disk gold electrodes Electrochemical measurements were performed using the BAS CV-100W potentiostat as above, however, in this case the reference electrode was an Hg|Hg2Cl2|KClsat electrode (SCE, 242 mV vs. NHE), and the counter electrode was a bare gold electrode. The supporting electrolyte was a 100 mM phosphate buffer at pH 5.0. The cyclic voltammograms were recoded from +140 to +1040 mV vs. NHE at three different scan rates: 10, 50, and 250 mV/s. For cleaning the gold electrodes, they were first immersed in freshly prepared Piranha solution (3:1 H2SO4 98%: 25% H2O2 30%) for 5 min. Next, the electrode surface of the working gold electrode was polished on DP-Suspension M. Pita et al. / Electrochemistry Communications 8 (2006) 747–753 and on alumina FF slurry (0.25 lm and 0.1 lm, Stuers, Copenhagen, Denmark), rinsed with Millipore water, and sonicated between and after polishing for 10 min. Then the electrode was electrochemically treated in 0.5 M sodium hydroxide (100 mV/s scan rate; between 60 and 1360 mV vs. NHE), in 0.5 M sulphuric acid (100 mV/s; between 60 and 1840 mV vs. NHE), and finally in 100 mM EDTA in 0.1 M phosphate buffer at pH 5.0 for 30 min at the constant applied potential of +790 mV vs. NHE. The electrode was kept then in 0.5 M sulphuric acid until use, when it was rinsed thoroughly with water. The gold counter electrode was cleaned by dipping for 5 min into Piranha solution and then rinsed with water thoroughly. For physical adsorption of laccase on the electrode surface, the electrode was mounted with its surface facing up and 10 lL of the laccase solutions (approximately 10 mg/ ml of C. maxima, 8 mg/ml of T. ochracea, and 18 mg/ml of T. hirsuta) were placed on the gold surface. The electrode was covered to avoid evaporation, and let it to react for 3 h. Covalent binding of laccase onto the electrode was accomplished through different self-assembled monolayers (SAMs) interfaces. First, gold electrodes were immersed into a 100 mM thiol solution. For L-cysteine and cystamine, the solvent for the thiol was Milli-Q water, and for 4-amino-thiophenol the solvent was absolute ethanol. The SAM was let to assemble for at least 4 h. In parallel, 10 lL of the laccase solution (same concentrations as taken for physical absorption, see above) were transferred into 0.2 ml of a 10 mg/ml NaIO4 in H2O at pH 5.0 and react for 30 min (pH is given by the periodate salt), in order to oxidise the sugar residues on the enzyme surface. After that, the pH was raised to 7 by adding 0.4 ml of 100 mM phosphate buffer. The oxidised sugar residues were allowed to react with the primary amine functionality present in the SAM, forming a Schiff base, after immersing the waterrinsed SAM-modified electrode into the modified enzyme solution for 1 h. Afterwards, the electrodes were ready to be used. 2.4. Measurements of H2O2 production The concentrations of H2O2 were measured spectrophotometrically using Uvikon 930 spectrophotometer (Kontron Instrument, Everett, MA, USA). A 0.89 ml aliquot was taken from the electrochemical cell and placed into a 1 ml cuvette. Then, ABTS (9.1 mM) and HRP (0.5 mg/ ml) were added as electron donor and catalyst, respectively, in order to get a final volume of 1 ml. The concentration of oxidised ABTS was measured at 405 nm (e = 36,800 M1 cm1), which corresponds to the concentration of H2O2 according to the following reaction: HRP H2 O2 þ ½ABTSred þ 2Hþ ! 2H2 O þ ½ABTSox ð1Þ Control experiments (measurements without applying any potential on the electrode) were performed and the results 749 were taken into account for the interpretation of the results. 3. Results and discussion Recently, ET between a bare gold electrode and the T1 site of T. hirsuta laccase in solution was undisputedly confirmed by spectroelectrochemical measurements [35]. It was shown that the blue colour vanishes when the applied potential was switched from being oxidative to being reductive. The bleaching was associated with the reduction of the T1 site of T. hirsuta laccase in the gold capillary electrode under anaerobic conditions [35]. In this work similar results were obtained for two other fungal laccases (C. maxima (Fig. 1) and T. ochracea (data not shown)). The redox transformations of the enzymes were found to be reversible and the laccases could be re-oxidised, either by applying an oxidising potential or by the introduction of oxygen. In our former work [35], only redox transformations of the T1 site were studied spectroelectrochemically; in the present work the redox state of both the T1 and T3 copper sites have been controlled simultaneously with a broader spectral measurements (330 nm region – T3 copper site, 600 nm region – T1 copper site). The possibility to obtain fully reduced enzymes was shown for all three enzymes (e.g., Fig. 1, curve 3 for C. maxima laccase). It should be mentioned that more than 70% of the initial enzyme activity is retained after the DET electrochemical titration procedure from oxidised to reduced enzyme and vice versa. The activity of the laccases was measured before and after redox transformations using ABTS as enzyme substrate [35]. In spite of the retained activity of the enzymes inside the capillary gold electrode and the possibility for DET between the electrode and the enzymes, only small bioelectrocatalytic currents were obtained in the presence of molecular oxygen (see Fig. 1, inset for C. maxima laccase). The electrochemistry of T. hirsuta, T. ochracea, and C. maxima laccases inside the capillary gold electrodes was further investigated with cyclic voltammetry. For all studied enzymes the anodic wave revealed irreversible electrochemistry; after the electrode had been subjected to an oxidative potential, the anodic wave became much less pronounced. The difference between the anodic peak potentials was rather small and found to be about 30 mV for the three laccases. Our results indicate that the oxidation process of the different copper ions in the active sites of laccases is significantly faster than that of the reduction process. These data are in good agreement with previously found behaviours of plant and fungal laccases on different gold electrodes [32,33,35,37]. However, it should be noticed that the electrochemical behaviour of C. maxima and T. ochracea laccases inside the capillary gold electrode was different from the behaviour shown by the enzyme from T. hirsuta [35]. Mainly, a much less pronounced (T. ochracea laccase) or even the absence (C. maxima) of a cathodic Faradaic process was observed, which suggested a sufficiently low 750 M. Pita et al. / Electrochemistry Communications 8 (2006) 747–753 1.6 3 E p ,a = 560 mV 1 1.35 2 Current / μA 0 Absorbance 1.1 -3 0.85 T3 -6 0.6 0 200 400 600 800 1000 Potential / mV vs. NHE 1 0.35 T1 2 3 0.1 300 350 400 450 500 550 600 650 700 750 800 850 Wavelength / nm Fig. 1. Electrochemical reactions of Cerrena maxima laccase inside the capillary gold electrode (0.1 M phosphate buffer, pH 6.5; the enzyme concentration was approximately 8 mg/ml). Mediatorless spectroelectrochemical oxidation and reduction of the laccase; 1, 2, and 3 are absorbance spectra of the enzyme at 875 mV (‘‘fully oxidised’’), 300 mV (‘‘partly reduced’’), and 200 mV (‘‘fully reduced’’), respectively. Inset: electroreduction of oxygen (second scan) inside the capillary gold electrode with (1) and without (2) Cerrena maxima laccase (scan rate – 10 mV/s; start potential – 1000 mV. rate constant for the heterogeneous ET compared to the value for T. hirsuta laccase (0.7 · 104 cm s1) calculated in [35]. To study the behaviour of the immobilized enzyme, cyclic voltammetry measurements of gold disc electrodes modified with the laccases have been performed (data not shown). The CVs obtained under anaerobic conditions for T. ochracea laccase contained only one single high potential ET process at +857 mV (vs. NHE). The process is more clearly pronounced when the scan rate was high. Similar CVs were obtained for C. maxima laccase, whereas the CVs of T. hirsuta laccase contained two well-pronounced ET processes at +486 and +903 mV vs. NHE. It should be noticed that the shape of the CVs for laccases adsorbed on bare gold electrodes strongly depends on the time allowed for the adsorption of the enzymes as well as the concentrations of the enzymes. For instance, when a low concentration of T. hirsuta laccase was used, only a weak high redox potential process at approximately +900 mV was observed, whereas in the case of a high concentration of the enzyme (18 mg/ml) and 1 h of adsorption a well-pronounced low-redox potential process could be clearly seen (data not shown). Most likely, both the adsorption time and the enzyme concentration strongly affect the orientation of the laccase on the surface along with partial denaturation of the enzyme on the electrode. A strong Faradaic current due to catalytic oxygen reduction under aerobic condition is observed in Fig. 2A (curve 1), starting at a potential of approximately +320 mV vs. NHE. It is also seen that addition of sodium fluoride to the solution does not affect this process. The conclusion drawn from this fact is that fluoride ions, a strong inhibitor of fungal and plant laccases, does not inhibit oxygen reduction on gold electrodes modified with adsorbed laccases. It can be suggested that the oxygen bioelectroreduction at laccase/gold electrodes takes place with no involvement of the intramolecular ET, which is hindered by fluoride ions [10,11,38]. The electrochemistry of laccases immobilised on a gold surface through covalent attachment to an amine-terminated-self assembled monolayer has also been studied for T. hirsuta, T. ochracea, and C. maxima laccases. Three different thiols have been used in the present study, viz. cystamine, L-cysteine, and 4-aminothiophenol. A well-pronounced and stable electrochemical signal was observed only in the case of 4-aminothiophenol-modified gold electrodes. It was shown that the electrochemistry of the enzymes on the modified thiol electrodes under anaerobic conditions resulted in two well-pronounced ET processes, at low and high potentials, with an Em5 of approximately 400 and 800 mV, respectively (data not shown). The peak current for the reduction process is lower than that for M. Pita et al. / Electrochemistry Communications 8 (2006) 747–753 A 2 0 4 Current/ µA -1 -2 -3 1 H 2O 2 Aerobic (1) Anaerobic (2) NaF (3) NaF anaerobic (4) -4 3 -5 200 400 600 800 1000 Potential / mV vs. NHE B 1 0 Current/ µA -1 4 2 -2 -3 H 2O 2 3 -4 Aerobic anaerobic NaF NaF anae -5 -6 H 2O 1 -7 200 400 600 800 (1) (2) (3) (4) 1000 Potential / mV vs. NHE Fig. 2. Representative cyclic voltammograms of (A) Cerrena maxima and (B) Trametes hirsuta laccases immobilised on (A) bare and (B) 4aminothiophenol self-assembled monolayer (SAM) modified gold (BAS) disc electrodes (0.1 M phosphate buffer pH 5.0; scan rate – 10 mV/s; second scan). (A) CVs of Cerrena maxima laccase under aerobic and anaerobic conditions, and in the absence or presence of sodium fluoride starting at 1040 mV. (B) CVs of Trametes hirsuta laccase under aerobic or anaerobic conditions, and in the absence or presence of sodium fluoride starting at 1040 mV. the oxidation. These features of the CVs are similar to those CVs of the enzyme on bare gold electrodes despite the fact that a self-assembled monolayer used here usually avoid enzyme denaturation on metal electrode surfaces. Taking into account this asymmetry between the reduction and oxidation currents, which has also been shown previously for plant laccases [37], it can be suggested that the difficulties with electroreduction of laccases related to the kinetics of the redox center reaction, which is lower than the oxidation of the electrodes, is due to the slow intramolecular ET process of the enzymes. In Fig. 2B, curve 1, a bioelectroreduction of molecular oxygen by T. hirsuta laccase immobilised on the thiol-modified gold electrodes is clearly seen. The electrochemical signal starts slightly from +625 mV vs. NHE, however, a more drastic increase is 751 seen only from +510 mV vs. NHE. Here it can be mentioned that, to our best knowledge, this potential is the highest potential for bioelectrocatalytic reduction of oxygen by laccase on a gold electrode reported up today. In other words, this is the closest value to the redox potential of the T1 site of the enzyme, which is found to be of 780 mV vs. NHE [13,35]. This CV is compared with another CV registered in the presence of sodium fluoride, and for this CV it only appears the reduction wave analogous to the wave presented for laccase-modified gold electrodes without thiols (cf. Fig. 2A and B, curves 3). Also, in the absence of oxygen, there is no biocatalytic response on the electrode (curves 2 and 4). A proposed explanation for our results could be that, in the presence of fluoride ions or when the enzyme is adsorbed on bare gold, instead of reduction of oxygen to water (natural laccase activity), oxygen is bioelectrocatalytically reduced to hydrogen peroxoxide at a much lower potential. In order to prove this hypothesis the possible production of hydrogen peroxide by T. hirsuta laccasemodified electrode have been measured via ABTS oxidation catalysed by HRP, either T. hirsuta is adsorbed on bare gold or covalently attached to a 4-aminothiophenol SAM-modified electrode. The results are shown in Fig. 3. In the case of bare gold electrodes (Fig. 3A) it is clearly seen that when a potential of +440 mV was applied on the electrode, only a residual production of hydrogen peroxide was observed. However, when the potential was lowered to +240 mV vs. NHE, a bare gold electrode catalyses the production of a small amount of H2O2, while the laccase modified bare gold electrode catalyses a 3 times higher production of H2O2 (from 12 nmol for a bare electrode to 36 nmol for a laccase-modified bare gold electrode per 70 min). When Trametes hirsuta laccase was immobilised on a self-assembled monolayer, the production of H2O2 is the same with or without the enzyme at a high potential (+440 mV; Fig. 3C), and is equal to the production by the enzyme-modified electrode when the electrode was polarised at a low potential (+240 mV; 40 nmol). The interesting result was that, when applying +240 mV to the SAM modified electrode without laccase, the H2O2 production was 70 times higher (2800 nmol) compared to the laccase-modified SAM-gold electrode (Fig. 3B). The covalent attachment of the laccase reduces drastically this effect because the enzyme reduces the oxygen and there is almost no oxygen left at the gold electrode surface that could be reduced to form hydrogen peroxide. In conclusion, bioelectroreduction of oxygen has been obtained from three different laccases when immobilised on gold electrodes. Immobilisation on bare gold electrodes gives a biocatalytic reduction of molecular oxygen, but this does not occur along the natural enzyme reduction pathway as long as the reduction product is hydrogen peroxide instead of water, and remains insensitive to sodium fluoride inhibition: Laccase O2 þ 2Hþ þ 2e ƒƒƒ! H2 O2 ð2Þ 752 M. Pita et al. / Electrochemistry Communications 8 (2006) 747–753 A 40 Bare 240 (1) Bare 440 (2) Lcs Bare 240 (3) Lcs Bare 440 (4) H2O2 / nmol 30 3 20 1 10 4 2 0 0 15 30 45 60 75 Time / min B C 3000 SAM 240 Lcs SAM 240 H2O2 / nmol H2O2 / nmol 2500 2000 1500 1000 40 SAM 440 Lcs SAM 440 30 20 10 500 0 0 0 15 30 45 60 75 0 Time / min 15 30 45 60 75 Time / min Fig. 3. Hydrogen peroxide production on bare and thiol-modified gold electrodes polarised at 240 mV and at 440 mV vs. NHE, with and without immobilised Trametes hirsuta laccase. (A) Bare gold electrode. (B and C) 4-aminothiophenol-modified gold electrode. This redox process probably does not take place using all the four copper ions, but only some of them. On the other hand, when the enzyme is immobilised on the electrode through the 4-aminothiophenol self-assembled monolayer, a mixed process of electroreduction of oxygen to water and to hydrogen peroxide appears. The electroreduction to water is due to the enzyme’s native activity, and the hydrogen peroxide production finds an explanation on whether a small amount of oxygen is reduced by 4-aminothiophenol or by a non-native activity of the enzymes: Laccase 2O2 þ 4Hþ þ 4e ƒƒƒ! 2H2 O ð3Þ Thus, the previously proposed schemes of the function of laccase on carbon and gold electrodes [2,35,39], where the enzyme is differently oriented onto the electrode surface, can also be used in order to explain differences in the electroactivity of laccases on bare and thiol-modified gold electrodes. Acknowledgements The authors thank Dr. Antonio López de Lacey for critical reading and helpful suggestions. The work has been financially supported by the Swedish Research Council, by the Swedish Institute (SI), by the Consejerı́a de Educación de la Comunidad de Madrid, and the European Social Funding (F.S.E.). The SI is acknowledged for the support of a postdoctoral fellowship for S.S. References [1] A.L. Ghindilis, P. Atanasov, E. Wilkins, Electroanalysis 9 (1997) 661. [2] S. Shleev, J. Tkac, A. Christenson, T. Ruzgas, A.I. Yaropolov, J.W. Whittaker, L. Gorton, Biosens. Bioelectron. 20 (2005) 2517. [3] A.I. Yaropolov, A.N. Kharybin, J. Emnéus, G. Marko-Varga, L. Gorton, Anal. Chim. Acta 308 (1995) 137. [4] R.S. Freire, N. Duran, L.T. Kubota, Talanta 54 (2001) 681. [5] B. Haghighi, L. Gorton, T. Ruzgas, L.J. Jönsson, Anal. Chim. Acta 487 (2003) 3. [6] A. Jarosz-Wilkolazka, T. Ruzgas, L. Gorton, Enzyme Microb. Technol. 35 (2004) 238. [7] S.C. Barton, J. Gallaway, P. Atanassov, Chem. Rev. 104 (2004) 4867. [8] S.C. Barton, H.-H. Kim, G. Binyamin, Y. Zhang, A. Heller, J. Am. Chem. Soc. 123 (2001) 5802. [9] S.C. Barton, M. Pickard, R. Vazquez-Duhalt, A. Heller, Biosens. Bioelectron. 17 (2002) 1071. [10] A.I. Yaropolov, O.V. Skorobogat’ko, S.S. Vartanov, S.D. Varfolomeyev, Appl. Biochem. Biotechnol. 49 (1994) 257. M. Pita et al. / Electrochemistry Communications 8 (2006) 747–753 [11] E.I. Solomon, U.M. Sundaram, T.E. Machonkin, Chem. Rev. 96 (1996) 2563. [12] A.A. Leontievsky, T. Vares, P. Lankinen, J.K. Shergill, N.N. Pozdnyakova, N.M. Myasoedova, N. Kalkkinen, L.A. Golovleva, R. Cammack, C.F. Thurston, A. Hatakka, FEMS Microbiol. Lett. 156 (1997) 9. [13] S.V. Shleev, O.V. Morozova, O.V. Nikitina, E.S. Gorshina, T.V. Rusinova, V.A. Serezhenkov, D.S. Burbaev, I.G. Gazaryan, A.I. Yaropolov, Biochimie 86 (2004) 693. [14] R.M. Wynn, H.K. Sarkar, R.A. Holwerda, D.B. Knaff, FEBS Lett. 156 (1983) 23. [15] K.-S. Shin, Y.-J. Lee, Arch. Biochem. Biophys. 384 (2000) 109. [16] B.R.M. Reinhammar, Biochim. Biophys. Acta 275 (1972) 245. [17] F. Xu, W. Shin, S.H. Brown, J.A. Wahleithner, U.M. Sundaram, E.I. Solomon, Biochim. Biophys. Acta 1292 (1996) 303. [18] P. Schneider, M.B. Caspersen, K. Mondorf, T. Halkier, L.K. Skov, P.R. Østergaard, K.M. Brown, S.H. Brown, F. Xu, Enzyme Microb. Technol. 25 (1999) 502. [19] A. Klonowska, C. Gaudin, A. Fournel, M. Asso, J. Le Petit, M. Giorgi, T. Tron, Eur. J. Biochem. 269 (2002) 6119. [20] B. Surma-Slusarska, J. Leks-Stepien, J. Wood Chem. Technol. 21 (2001) 361. [21] M. Balakshin, C.-L. Chen, J.S. Gratzl, A.G. Kirkman, H. Jakob, J. Mol. Catal. B: Enz. 16 (2001) 205. [22] M. Paice, R. Bourbonnais, S. Renaud, S. Labonte, G. Sacciadis, R. Berry, M. Amann, A. Candussio, R. Mueller, Prog. Biotechnol. 21 (2002) 203. [23] Y. Wong, J. Yu, Water Res. 33 (1999) 3512. [24] A.M. Mayer, R.C. Staples, Phytochemistry 60 (2002) 551. 753 [25] K. Kruus, Kemia - Kemi 27 (2000) 184. [26] R.C. Minussi, G.M. Pastore, N. Duran, Trend. Food Sci. Technol. 13 (2002) 205. [27] N. Duran, M.A. Rosa, A. D’Annibale, L. Gianfreda, Enzyme Microb. Technol. 31 (2002) 907. [28] M.R. Tarasevich, V.A. Bogdanovskaya, N.M. Zagudaeva, A.V. Kapustin, Russian J. Electrochem. 38 (2002) 335. [29] F. Barriere, Y. Ferry, D. Rochefort, D. Leech, Electrochem. Commun. 6 (2004) 237. [30] V. Soukharev, N. Mano, A. Heller, J. Am. Chem. Soc. 126 (2004) 8368. [31] I.V. Berezin, V.A. Bogdanovskaya, S.D. Varfolomeev, M.R. Tarasevich, A.I. Yaropolov, Dokl. Akad. Nauk SSSR 240 (1978) 615. [32] M. Gelo-Pujic, H.H. Kim, N.G. Butlin, G.T. Palmore, Appl. Environ. Microbiol. 65 (1999) 5515. [33] C.-J. Yoon, H.-H. Kim, J. Korean Electrochem. Soc. 7 (2004) 26. [34] G. Gupta, V. Rajendran, P. Atanassov, Electroanalysis 16 (2004) 1182. [35] S. Shleev, A. Christenson, V. Serezhenkov, D. Burbaev, A. Yaropolov, L. Gorton, T. Ruzgas, Biochem. J. 385 (2005) 745. [36] T. Larsson, A. Lindgren, T. Ruzgas, Bioelectrochemistry 53 (2001) 243. [37] D.L. Johnson, J.L. Thompson, S.M. Brinkmann, K.A. Schuller, L.L. Martin, Biochemistry 42 (2003) 10229. [38] A.I. Yaropolov, A.N. Kharybin, J. Emnéus, G. Marko-Varga, L. Gorton, Bioelectrochem. Bioenerg. 40 (1996) 49. [39] S. Shleev, A. Jarosz-Wilkolazka, A. Khalunina, O. Morozova, A. Yaropolov, T. Ruzgas, L. Gorton, Bioelectrochemistry 67 (2005) 115.