Amperometric detection of methanol with a methanol dehydrogenase modified electrode sensor

Journal of

Electroanalytical Chemistry Journal of Electroanalytical Chemistry 601 (2007) 125–131 www.elsevier.com/locate/jelechem

Amperometric detection of methanol with a methanol dehydrogenase modified electrode sensor Qinfeng Liu, Jon R. Kirchhoff

*

Department of Chemistry, The University of Toledo, 2801 W. Bancroft Street, Toledo, OH 43606, United States Received 6 July 2006; received in revised form 24 October 2006; accepted 30 October 2006 Available online 18 December 2006

Abstract An amperometric enzyme electrode was developed by immobilizing the quinoprotein methanol dehydrogenase from Methylobacterium extorquens AM1 onto a glassy carbon electrode. Ferrocene (FC), ferrocene carboxylic acid, N,N,N 0 ,N 0 -tetramethyl-1,4-phenylenediamine (TMPD), N,N-dimethyl-p-phenylenediamine (DMPD), phenazine methosulfate (PMeS) and Wurster blue (WB) were evaluated as electron-transfer mediators. DMPD, TMPD and Fc were found to be effective electron-transfer mediators for the enzyme with only ferrocene exhibiting a stable response in the presence of oxygen. The best response for the detection of methanol was achieved with a 100 lM suspension of ferrocene in 100 mM Tris/HCl buffer pH 9.0. The sensor utilized very small quantities of enzyme, and exhibited excellent reproducibility and stability with a detection limit of 0.5 lM (S/N = 3) and a linear range of 0.5–200 lM. Ó 2006 Elsevier B.V. All rights reserved. Keywords: Enzyme electrode; Amperometric biosensor; Methanol dehydrogenase; Pyrroloquinoline quinone; Methanol detection

1. Introduction Methanol is one of the most widely used organic solvents, especially in industrial and household products. It is also potentially valuable as an alternative automobile fuel [1]. However, methanol exposure via inhalation and skin absorption may lead to toxic effects from headaches to blindness with direct digestion even leading to death [2,3]. Therefore, facile analysis procedures are important for monitoring methanol levels in the environment, in alcoholic beverages and for clinical diagnostic measurements. Methanol determination is also required in some biological processes. For example, methylotropic yeasts such as P. pastors and H. polymorpha are emerging as commercially important hosts for the expression and production of recombinant heterogeneous proteins [4,5] because they afford several advantages over the traditional E. coli sys-

*

Corresponding author. Tel.: +1 419 530 1515; fax: +1 419 530 4033. E-mail address: [email protected] (J.R. Kirchhoff).

0022-0728/$ - see front matter Ó 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.jelechem.2006.10.039

tem. In order to maximize the recombinant protein yield in these systems, it is critical to carefully maintain the methanol concentration in the ferment below toxic levels to ensure normal yeast growth [6]. Many different instrumental methods are used for methanol determination, including GC-FID [7–9], HPLC with electrochemical [10], fluorescence [11] or UV–visible [12] detection and FTIR [13]. One simple alternative is the use of an amperometric biosensor, which uses an enzyme to recognize and oxidize methanol. The resulting current signal is then proportional to the methanol concentration and is useful for quantitation. Most enzymatic alcohol biosensors have been designed for ethanol detection with only a few developed for methanol detection [10,14,15]. These biosensors typically utilized either nicotinamide (NAD+) dependent alcohol dehydrogenases (ADH, EC1.1.1.1) or alcohol oxidases (AOD, EC1.1.3.13) as the detection enzyme. One challenge for ADH enzymes is that oxidation of NAD+ is hard to achieve at an electrode surface without applying a high over-potential [16]. Furthermore, NAD+ is a free-diffusive soluble cofactor and can tend to leak out of the immobilized enzyme and negatively affect sensor

126

Q. Liu, J.R. Kirchhoff / Journal of Electroanalytical Chemistry 601 (2007) 125–131

performance [17]. In the case of AOD enzymes, molecular O2 acts as the electron acceptor and the resulting biosensor is based either on monitoring the consumption of O2 or the generation of H2O2. Therefore, residual O2 in the sample solution will affect the current signal of AOD alcohol biosensors, while H2O2 can introduce interferences from biological species that are oxidized by H2O2. It is also difficult to find mediator systems for this type of sensor, because artificial mediators rarely can compete with O2, the native electron acceptor of AOD enzymes. A third class of alcohol dehydrogenases, pyrroloquinoline quinone (PQQ) dependent alcohol dehydrogenases (EC1.1.99.8), has also been employed in biosensor designs for ethanol [18–30]. In contrast, methanol dehydrogenase (MDH) from which PQQ was first purified [31] has been rarely used in sensing applications [27,32]. In one case, MDH was incorporated into a bioelectrochemical fuel cell for the enzymatic oxidation of methanol to formic acid to produce low levels of power output [32]. The coulometric detection of primary alcohols was also demonstrated using the MDH-based biofuel cell [32]. In a second example, a carbon paste electrode modified with MDH was also reported for the amperometric determination of ammonia [27]. Advantages of the PQQ alcohol dehydrogenases are that O2 and H2O2 are not involved in the catalytic reaction and the cofactor remains associated with the enzyme. One drawback to their use in electrochemical-based sensors is the difficulty for direct electron transfer between the enzymes and an electrode. The biological electron acceptor for MDH is a soluble cytochrome c [33–36], and when compared to small molecules, proteins are not good choices for electron-transfer mediators in sensor devices. Therefore, in this paper, we examine several mediators and their application for the development of a simple amperometric enzyme sensor based on MDH.

2. Experimental 2.1. Materials MDH was isolated from Methylobacterium extorquens AM1 (American Type Culture Collection, ATCC14718) by the method of Day and Anthony [37] and purified by the method of Liu et al. [38]. Phenazine methosulfate (PMeS), N,N,N 0 ,N 0 -tetramethyl-1,4-phenylenediamine (TMPD), N,N-dimethyl-p-phenylenediamine dihydrochloride (DMPD), ferrocene (Fc) and ferrocenecarboxylic acid (FcCOOH) were purchased from Aldrich. Wurster blue (WB) was prepared from TMPD according to a literature method [39]. HPLC grade methanol was obtained from Fisher Scientific. Cellulose semi-permeable membranes (10 kD cutoff, 20 lm thickness) were purchased from Fisher. All other reagents were analytical reagent grade and used without further purification. Aqueous solutions were prepared with distilled deionized water purified to a

resistivity of at least 17 MX-cm by a Barnstead B pure water purification system. 2.2. Instrumentation All electrochemical experiments were performed with a Bioanalytical Systems (BAS) 100B/W electrochemical analyzer using a conventional three-electrode electrochemical cell. A standard glassy carbon (MF-2012, BAS) or an enzyme modified electrode was used as the working electrode. A coiled platinum wire (MW-1033, BAS) and a Ag/AgCl (3 M NaCl) (MF-2052, BAS) electrode were employed as the auxiliary and reference electrodes, respectively. 2.3. Preparation of the MDH enzyme electrode The enzyme electrode was prepared by first polishing a glassy carbon electrode with alumina sanding gel (ultra micro fine, Nicsand) on a felt polishing cloth (Buehler). The polished electrode was then thoroughly rinsed with deionized water and methanol, and finally sonicated for 3 min in distilled water. MDH (0.3–1.0 units (6–10 lL)) from a 20 mM phosphate buffer pH 7.0 stock solution was pipetted onto the center of the glassy carbon surface and dried in air for 30 min. To cover the disk electrode surface with a uniform layer of enzyme solution 6–10 lL was sufficient. The electrode was then covered with a semipermeable cellulose membrane, which was secured with an O-ring. The MDH enzyme electrode was stored in dry conditions at 0 °C before use. 2.4. Electrochemical methods Evaluation of the different electron-transfer mediators for the MDH electrode was performed by cyclic voltammetry. The MDH sensor was characterized by amperometric detection of methanol in 100 mM Tris/HCl buffer pH 9.0 unless otherwise specified. The solution was purged with high purity Ar for 15 min and kept under Ar during the experiment for evaluation of PMeS, WB, TMPD or DMPD as mediators. Since Fc is only sparingly soluble in water, it was first dissolved in acetonitrile (10 mg/mL) and then suspended into Tris buffer. The final concentration if soluble would be equivalent to 100 lM. Amperometric experiments were performed at room temperature under hydrodynamic conditions induced by a magnetic stirring bar. The applied potential for amperometric detection for methanol varied depending on the mediator used. 3. Results and discussion 3.1. Evaluation of electron-transfer mediators for amperometric detection of methanol In the absence of a mediator, the MDH-based methanol sensor did not respond to methanol even with detection

Q. Liu, J.R. Kirchhoff / Journal of Electroanalytical Chemistry 601 (2007) 125–131 Table 1 Summary of the redox potentials for different mediators

20 nA

10 μM

0

60

127

20 μM

120

180

Mediator or cofactor

E° 0 , Redox potential measured in 100 mM Tris/HCl pH 9.0, mV vs. Ag/AgCl

Potential applied for alcohol detection, mV vs. Ag/AgCl

PQQa PMeS WB, TMPD DMPD Fc FcCOOH

260 160 50 110 160 320

– 0 150 200 250 340

50 μM

240

300

360

Time, s

a

Fig. 1. Amperometric response of a MDH enzyme electrode in 100 mM Tris/HCl pH 9.0 to repetitive methanol injections in the absence of a mediator. MDH (0.3 Units) was immobilized on the electrode. Detection potential: 0.6 V vs. Ag/AgCl.

potentials as high as 0.6–0.8 V vs. Ag/AgCl (Fig. 1). The structure of MDH reveals the PQQ cofactor is located ˚ away inside the protein shell of the enzyme more than 10 A from the surface [40]. Such isolation of the redox cofactor suggests a mediator molecule is necessary to act as an electron-transfer relay in our sensor scheme. Unmediated redox behavior was observed for PQQ fructose dehydrogenase [41] and the quinohemoprotein alcohol dehydrogenase, which contains additional heme groups closer to the protein surface that facilitate direct electron-transfer [24]. One elegant approach to facilitate electron-transfer between the enzyme and an electrode was demonstrated by Willner and co-workers in the development of a glucose sensing scheme [42]. In this case the FAD cofactor of glucose oxidase was linked to ferrocene or tethered to an electrode surface through a PQQ electron-transfer relay linker. Upon reconstitution of the apoenzyme with FAD, the electron-transfer mediators were in close proximity to the cofactor, therefore decreasing the electron-transfer distance and enabling observation of a measurable current response for conversion of glucose to gluconic acid. Given the simple mode of entrapment of MDH on the electrode surface, addition of a mediator to the analysis solution was utilized. PMeS and WB are known to oxidize the reduced enzyme and are used in the enzyme activity assay for MDH [43]. Both were evaluated as mediators for the development of the amperometric enzyme sensor along with DMPD, FcCOOH, Fc and TMPD, which is the reduced form of WB. The redox potentials for each molecule in 100 mM Tris/HCl buffer pH 9.0 were determined by cyclic voltammetry and are summarized in Table 1. Based on the relative redox potentials of the mediator to PQQ, it is thermodynamically feasible for each molecule to act as a mediator for MDH and ensure regeneration or oxidation of the reduced enzyme. It is advantageous that the mediator redox potential is not too positive in order to minimize the background response and its effect on the electron-transfer process [39,45]. Once incorporated into the sensor a potential slightly more positive than the oxidation peak potential of the mediator was applied to the

Redox potential of PQQ at pH 9.0 was obtained from the literature [44].

MDH electrode to evaluate the sensor system for detection of methanol. The detection potentials are also summarized in Table 1. The response of the MDH electrode greatly depended on the mediator used. Fig. 2a depicts the amperometric response of the MDH electrode to repetitive injections of methanol using 500 lM WB as the mediator. The electrode showed very good sensitivity for methanol, however, at

0.2 μA

10 μM MeOH each injection

0

400

800

Stirring disturbance

0.2 μA

20 μM

1200

1600

100 μM 50 μM

10 μM

5 μM 10 μM

0

400

800

1200

1600

Time, s Fig. 2. Effect of NH4Cl on the sensor performance in (a) 100 mM Tris/ HCl pH 9.0 with 500 lM WB as the mediator, 10 lM injections of methanol; (b) same as (a) with 15 mM NH4Cl. MDH (0.3 Units) was immobilized on the electrode.

128

Q. Liu, J.R. Kirchhoff / Journal of Electroanalytical Chemistry 601 (2007) 125–131

higher concentrations of methanol the electrode became saturated and stopped functioning. In an attempt to improve the sensitivity even further NH4Cl, which was reported to efficiently activate MDH in the spectroscopic enzyme activity assay of MDH when PMeS or WB was used as the artificial electron acceptor [33], was added. For this application, NH4Cl accelerated the deterioration of the electrode instead of providing an activation effect (Fig. 2b) perhaps indicating differences in the assayed reaction between these two methods. PMeS, WB and TMPD were sensitive to light and oxygen. The absorbance of a solution of WB dissolved in 100 mM Tris/HCl buffer pH 9.0 decreased by more than 50% in 45 min, while TMPD gradually obtained the blue color of WB within a few hours in aerobic conditions in the same buffer. Purging the solution with argon suppressed the color change indicating the oxidation of TMPD by dissolved oxygen. PMeS stock solutions were stored in the dark, otherwise the solution gradually changed color within hours. TMPD and DMPD effectively mediated the amperometric detection of methanol and exhibited catalytic oxidative currents for methanol (Figs. 3 and 4). TMPD (Fig. 3b) exhibited a higher ratio of catalytic current (ipa,2/ipa,1  5.2) compared to DMPD (Fig. 4b; ipa,2/ipa,1  1.2) as well as a greater degree of reversibility (TMPD, DEp1 = 55 mV, ipa,1/ipc,1  1; DMPD, DEp1 = 57 mV, ipa,1/ipc,1  4). In comparison to PMeS and WB, the addition of NH4Cl had only a slight effect on the electrode response. With TMPD, methanol concentrations as low as 1 lM were detected. The main disadvantage is the need to maintain anaerobic conditions to minimize mediator oxidation. To the best of our knowledge DMPD has never been used as an electron-transfer mediator for quinoprotein dehydrogenases or other oxido-reductases. The electron-transfer pathway for MDH is believed to be two one-electron transfers between the enzyme and a one-electron acceptor such as cytochrome cl, PMeS or WB [46], while the electrochemical oxidation of DMPD was usually observed as a twoelectron reversible process [47]. Recently the electrochemical oxidation of DMPD was described as one fast electron transfer (102 times faster) separated from a second slower electron transfer through ultrasound treatment with a cation radical intermediate similar to WB [48]. Fig. 4 illustrates that the electron-transfer between DMPD and MDH must be fast enough to compete with the second step of the electrode oxidation of DMPD. A shoulder was also observed in cyclic voltammogram (2) in Fig. 4b, which may be due to the separation of the two one-electron transfer processes by the catalytic reaction between the methanol– MDH complex and DMPD+. FcCOOH was also reported to mediate the electrontransfer of PQQ dependent MDH in cyclic voltammetric studies [49]. In the presence of FcCOOH, the MDH electrode exhibited a rapidly decreasing baseline and irreproducible response to methanol. Methylene blue and hexacyanoferrate were also evaluated, but neither facili-

0.5 μA

1 μM MeOH injection 2.5 μM MeOH 10 μM MeOH

0

500

1000

1500

2000

Time, s

(1)

0.4 μA

(2)

300

100

-100

Potential, mV Fig. 3. Amperometric response of the MDH electrode for (a) 100 lM TMPD in 100 mM Tris/HCl pH 9.0 with 15 mM NH4Cl and (b) cyclic voltammetric responses without (1) and with (2) 195 lM methanol. Scan rate is 50 mV/s. MDH (0.3 Units) was immobilized on the electrode. Inset: expanded version of 0–500 s.

tated the electron-transfer process in our electrochemical scheme. An unexpected observation was that a saturated solution of Fc was a very good mediator for MDH although it is only slightly soluble in aqueous solution (Fig. 5). This is supported by the voltammograms in Fig. 5b where the reversible one-electron oxidation of Fc was observed along with a significant catalytic response. Poor solubility of Fc compared to the oxidized form, Fc+, resulted in some adsorption of Fc on the electrode surface. This is indicated in voltammogram (1) by a DEp1  39 mV and a current ratio of ipc,1/ipa,1  0.27, both of which deviate from the theoretical values of DEp = 59 mV and ipc/ipa  1.0 for a reversible one-electron transfer. Upon addition of

Q. Liu, J.R. Kirchhoff / Journal of Electroanalytical Chemistry 601 (2007) 125–131

129

0.2 μA

10 μM

10 μM MeOH injection each

5 μM MeOH injection

0.2 μA

0

500

1000

1500

2000

Time, s 0

400

800

1200

1600 (1)

Time, s

*

(1)

0.5 μA

*

(2)

400

(2) 1 μA

300

200

0

Potential, mV

100

-100

Fig. 5. Amperometric response of the MDH electrode for (a) the equivalent of 100 lM Fc suspended in 100 mM Tris/HCl pH 9.0 with 15 mM NH4Cl and (b) cyclic voltammetric responses without (1) and with (2) 160 lM methanol. Scan rate is 50 mV/s. MDH (0.3 Units) was immobilized on the electrode.

Potential, mV Fig. 4. Amperometric response of the MDH electrode for (a) 100 lM DMPD in 100 mM Tris/HCl pH 9.0 with 15 mM NH4Cl and (b) cyclic voltammetric responses without (1) and with (2) 125 lM methanol. Scan rate is 50 mV/s. MDH (0.3 Units) was immobilized on the electrode. * denotes appearance of a shoulder.

160 lM methanol (Fig. 5(b), voltammogram 2), the oxidative peak current was enhanced indicating the catalytic nature of the response (ipa,2/ipa,1  4.0, ipc,2/ipc,1  6.5). The MDH enzyme electrode with Fc as the mediator was very stable, exhibited a highly reproducible response and was not affected by the presence of oxygen or NH4Cl compared to many of the other mediators. For these reasons Fc was selected as the mediator for the development and complete characterization of the MDH-based amperometric methanol sensor. 3.2. Optimization and evaluation of the MDH enzyme electrode for the detection of methanol Beside the electron-transfer mediator, the solution pH, the amount of enzyme used and the enzyme specific activity were optimized prior to characterization of the MDH electrode. Comparative cyclic voltammograms with Fc as the

mediator demonstrated that as the pH was increased from pH 7.0 (100 mM phosphate) to 9.0 (100 mM Tris/HCl) the catalytic oxidation current of methanol at the MDH electrode increased by approximately a factor of two (data not shown). This result is consistent with the behavior of other quinoprotein alcohol dehydrogenase enzymes [50]. Above pH 9, MDH begins to denature. Therefore, 100 mM Tris/HCl pH 9 was used for all quantitative evaluations of the electrode. Fig. 6 depicts the response of the MDH electrode as a function of methanol concentration for two electrodes with a constant enzyme amount (0.6 units), but with specific activities of 0.5 units/mg (1) and 5.4 units/mg (2). The response to methanol exhibited improvements in sensitivity and the linear detection range with enzyme of higher specific activity (2). Therefore, enzyme of the highest purity was prepared from the crude extract and incorporated into all electrodes. This is very important since the amount of enzyme that can be used is limited due to the small size of the working electrode. Typically 6–10 lL of purified enzyme was applied to construct the sensor. On the other hand, the small amount of enzyme is a clear advantage of this sensor compare to coulometric detection scheme used with the MDH biofuel cell [32].

130

Q. Liu, J.R. Kirchhoff / Journal of Electroanalytical Chemistry 601 (2007) 125–131 0.50

4.50

0.8

0.40

(2)

0.30

4.00

Sensor Response (i-i0), µA

0.20 y = 0.0091x 0.10

Sensor Response (i-i 0), µA

3.50

2

R = 0.9806

0.00 05

10 15 20 25

y = 0.0194x

3.00

2

R = 0.9999

2.50 2.00 1.50

y = 0.0090x R2 = 0.9976 0.6

y = 0.0084x R2 = 0.9933

0.4

0.2

(1) 0

1.00

0

25

50

75

100

[CH3OH], μM

0.50 0.00 0

50

100

150

200

250

300

[MeOH], μM Fig. 6. Plot of the amperometric response of the MDH electrode for the immobilization of MDH of different specific activity in 100 mM Tris/HCl pH 9.0, 100 lM TMPD. (j) 0.6 units MDH, 5.4 units/mg and (s) 0.6 units MDH, 0.5 units/mg. The inset shows an expanded view of the data at low methanol concentrations.

Fig. 7. Comparative response of the MDH electrode as a function of time in 100 mM Tris/HCl pH 9.0 with the equivalent of 100 lM Fc suspended in solution. (m) freshly made electrode with 0.3 units MDH and (s) after one week of storage in a desiccator at 0 °C. Repetitive 5 lM injections of methanol.

Table 2 Performance characteristics of the MDH electrode MDHa Limit of detection, S/N = 3

3.3. Sensor stability When Fc was used as the electron-transfer mediator, MDH showed very good stability in the sensor. Three calibration plots were obtained from measurements using a freshly prepared MDH electrode over the course of an eight hour time period (immediately after preparation, at 1.5 h and at 8 h). Between calibration intervals the electrode was rinsed and stored in Tris/HCl buffer and then reused for the next calibration measurement. There was no obvious loss of sensitivity for the sensor over this time period. The only difference occurred in the calibration plot at 8 h where the upper limit of linearity was slightly reduced. Repeat measurements over an extended time period of one week demonstrated that the electrode maintained its level of performance when tested after being stored in a desiccator at 0 °C. Deviations of less than 7% (Fig. 7) were observed.

Sensitivity comparisonb iMeOH/iEtOH Linear detection range Response time a b

0.5 lM MeOH 1 lM EtOH 1.4 0.5–200 lM MeOH 1–100 lM EtOH 10 s for MeOH and EtOH

0.6 units of MDH were used. Within linear ranges of both methanol and ethanol.

nol could limit the electrode selectivity for the detection of methanol in the presence of ethanol because of the nonspecificity of MDH [33]. The MDH electrode had a rapid response with similar linear ranges to both methanol and ethanol. The detection limit for methanol was 0.5 lM (S/N = 3) and for ethanol was 1 lM (S/N = 3). In comparison, the detection limit for ethanol at a carbon electrode modified with Type III ethanol dehydrogenase is reported as 1–5 lM (S/N = 2) [21]. 4. Conclusions

3.4. Sensor performance characteristics At the optimized conditions, the detection limit, linear range, and sensitivity of the MDH electrode to methanol were determined and are listed in Table 2. Since ethanol is the common interference for most methanol analyses, the MDH electrode response to ethanol was examined and included in Table 2. The current signal ratio of methanol compared to ethanol was 1.4. Although the relative current favors detection of methanol, the presence of etha-

An amperometric biosensor was constructed by immobilizing MDH, a PQQ dependent quinoprotein from Methylobacterium extorquens AM1, onto a glassy carbon electrode surface by semi-permeable membrane entrapment. The sensor utilizes very small quantities of enzyme, and exhibits excellent reproducibility and stability with the proper choice of electron-transfer mediator. DMPD, TMPD and Fc were found to be effective electron-transfer mediators for the detection of methanol in combination

Q. Liu, J.R. Kirchhoff / Journal of Electroanalytical Chemistry 601 (2007) 125–131

with the simple electrode design. However, due to the oxygen sensitivity of TMPD and DMPD, the MDH electrode was optimized using a suspension of Fc to create a sensor with a detection limit for methanol of 0.5 lM with a linear range up to 200 lM in 100 mM Tris/HCl buffer pH 9.0. Further efforts to improve the selectivity toward methanol compared to other primary alcohols are currently in progress. Acknowledgements Financial support from The University of Toledo College of Arts and Sciences for funding and operation of the Arts and Sciences Instrumentation Center is gratefully acknowledged. References [1] N.D. Brinkman, E.E. Ecklund, R.J. Nichols, Fuel Methanol: A Decade of Progress, Society of Automotive Engineers, Warrendale, PA, 1990. [2] Occupational Exposure to Methyl Alcohol, Division of Criteria Documentation and Standards Development, Department of Health, Education, and Welfare, Public Health Service, Center for Disease Control, National Institute for Occupational Safety and Health, Washington, DC, 1976. [3] Methanol Toxicity [microfilm], in: US Department of Health & Human Services, Public Health Service, Agency for Toxic Substances and Disease Registry, Atlanta, GA, 1992. [4] P.E. Sudbery, Curr. Opin. Biotechnol. 7 (1996) 517–524. [5] J.L. Cereghino, J.M. Cregg, FEMS Microbiol. Rev. 24 (2000) 45–66. [6] G. Gellissen, C.P. Hollenberg, Z.A. Janowicz, in: A. Smith (Ed.), Gene Expression in Recombinant Microorganisms, Marcel Dekker, New York, 1995, pp. 195–239. [7] P.A. de Paula Pereira, E.T. Sousa Santos, T. de Freitas Ferreira, J.B. de Andrade, Talanta 49 (1999) 245–252. [8] X.P. Lee, T. Kumazawa, K. Kondo, K. Sato, O. Suzuki, J. Chromatogr. B 734 (1999) 155–162. [9] J.F. Livesey, S.L. Perkins, N.E. Tokessy, M.J. Maddock, Clin. Chem. 41 (1995) 300–305. [10] F. Tagliaro, G. Schiavon, R. Dorizzi, M. Marigo, J. Chromatogr.Biomed. Appl. 563 (1991) 11–21. [11] A.I. HajYehia, L.Z. Benet, J. Chromatogr. A 724 (1996) 107–115. [12] S.H. Chen, H.L. Wu, C.H. Yen, S.M. Wu, S.J. Lin, H.S. Kou, J. Chromatogr. A 799 (1998) 93–99. [13] J.M. Garrigues, A. Perez-Ponce, S. Garrigues, M. de la Guardia, Vib. Spectrosc. 15 (1997) 219–228. [14] M. Blanco, J. Coello, H. Iturriaga, S. Maspoch, M. Porcel, Anal. Chim. Acta 398 (1999) 83–92. [15] C. Almuzara, O. Cos, M. Baeza, D. Gabriel, F. Valero, Biotechnol. Lett. 24 (2002) 413–417. [16] Z. Samec, P.J. Elving, J. Electroanal. Chem. 144 (1983) 217–234. [17] K. Habermu¨ller, M. Mosback, W. Schuhmann, J. Anal. Chem. 366 (2000) 560–568. [18] J. Razumiene, V. Gureviciene, V. Laurinavicius, J.V. Grazulevicius, Sens. Actuators B Chem. 78 (2001) 243–248. [19] W. Schuhmann, H. Zimmermann, K.V. Habermu¨ller, V. Laurinavicius, Faraday Discuss. 116 (2000) 245–255.

131

[20] M. Niculescu, T. Erichsen, V. Sukharev, Z. Kerenyi, E. Csoregi, W. Schuhmann, Anal. Chim. Acta 463 (2002) 39–51. [21] J. Razumiene, M. Niculescu, A. Ramanavicius, V. Laurinavicius, E. Csoregi, Electroanalysis 14 (2002) 43–49. [22] T. Ikeda, K. Kato, M. Maeda, H. Tatsumi, K. Kano, K. Matsushita, J. Electroanal. Chem. 430 (1997) 197–204. [23] S. Zhao, R.B. Lennox, Anal. Chem. 63 (1991) 1174–1178. [24] A. Ramanavicius, K. Habermu¨ller, E. Csoregi, V. Laurinavicius, W. Schuhmann, Anal. Chem. 71 (1999) 3581–3586. [25] V. Laurinavicius, J. Razumiene, B. Kurtinaitiene, I. Lapenaite, I. Bachmatova, L. Marcinkeviciene, R. Meskys, A. Ramanavicius, Bioelectrochemistry 55 (2002) 29–32. [26] M.E. Argall, G.D. Smith, Biochem. Mol. Biol. Int. 30 (1993) 491–497. [27] S.I. Suye, Y. Nakamura, S. Inuta, T. Ikeda, M. Senda, Biosens. Bioelectron. 11 (1996) 529–534. [28] E.C.A. Stigter, G.A.H. deJong, J.A. Jongejan, J.A. Duine, J.P. vanderLugt, W.A.C. Somers, Enzyme Microb. Technol. 18 (1996) 489–494. [29] E.C.A. Stigter, G.A.H. deJong, J.A. Jongejan, H.A. Duine, J.P. vanderLugt, W.A.C. Somers, J. Chem. Technol. Biotechnol. 68 (1997) 110–116. [30] T. Ikeda, D. Kobayashi, F. Matsushita, T. Sagara, K. Niki, J. Electroanal. Chem. 361 (1993) 221–228. [31] J.A. Duine, J. Frank, P.E.J. Verviel, Eur. J. Biochem. 108 (1980) 187– 192. [32] G. Davis, H.A.O. Hill, W.J. Aston, I.J. Higgins, A.P.F. Turner, Enzyme Microb. Technol. 5 (1983) 383–388. [33] C. Anthony, Adv. Microb. Physiol. 27 (1986) 113–210. [34] K.A.O. Matsushita, in: V.L. Davidson (Ed.), Principles and Applications of Quinoproteins, Marcel Dekker, New York, 1993, p. Ch. 3. [35] Y. Ohshiro, S. Itoh, in: V.L. Davidson (Ed.), Principles and Applications of Quinoproteins, Marcel Dekker, New York, 1993, pp. 309–330. [36] C. Anthony, in: V.L. Davidson (Ed.), Principles and Applications of Quinoproteins, Marcel Dekker, New York, 1993 (Chapter 2). [37] D.J. Day, C. Anthony, in: M.E. Lidstrom (Ed.), Methods in Enzymology, vol. 188, Academic Press, San Diego, CA, 1990, pp. 210–216. [38] Q. Liu, J.R. Kirchhoff, C.R. Faehnle, R.E. Viola, R.A. Hudson, Protein Exp. Purif. 46 (2006) 316–320. [39] L. Michaelis, S. Granick, J. Am. Chem. Soc. 65 (1943) 1747–1755. [40] M. Ghosh, C. Anthony, K. Harlos, M.G. Goodwin, C. Blake, Structure 3 (1995) 177–187. [41] J. Parellada, E. Domı´nguez, V.M. Ferna´ndez, Anal. Chim. Acta 330 (1996) 71–77. [42] E. Katz, A. Riklin, V. Heleg-Shabtai, I. Willner, A.F. Bu¨ckmann, Anal. Chim. Acta 385 (1999) 45–58. [43] J.A. Duine J. Frank, J. Westerling, Biochim. Biophys. Acta 524 (1978) 277–287. [44] K. Kano, K. Mori, B. Uno, T. Kubota, T. Ikeda, M. Senda, Bioelectrochem. Bioenerg. 23 (1990) 227–238. [45] R.A. Marcus, N. Sutin, Biochim. Biophys. Acta 811 (1985) 265–322. [46] J. Frank, M. Dijkstra, C. Balny, P.E.J. Verwiel, J.A. Duine, Antonie Van Leeuwenhoek, J. Microbiol. 56 (1989) 25–34. [47] A.D. Modestov, J. Gun, I. Savotine, O. Lev, J. Electroanal. Chem. 565 (2004) 7–19. [48] C.E. Banks, N.S. Lawrence, R.G. Compton, Electroanalysis 15 (2003) 243–248. [49] A.E.G. Cass, G. Davis, M.J. Green, H.A.O. Hill, J. Electroanal. Chem. 190 (1985) 117–127. [50] H. Gorisch, M. Rupp, Antonie Van Leeuwenhoek, J. Microbiol. 56 (1989) 35–45.