Catalytic activity of iron hexacyanoosmate(II) towards hydrogen peroxide and nicotinamide adenine dinucleotide and its use in amperometric biosensors

Analytica Chimica Acta 599 (2007) 287–293

Catalytic activity of iron hexacyanoosmate(II) towards hydrogen peroxide and nicotinamide adenine dinucleotide and its use in amperometric biosensors Petr Kotzian a , Tereza Jank˚u a , Kurt Kalcher b , Karel Vytˇras a,∗ a

Department of Analytical Chemistry, University of Pardubice, Nam. Cs. Legii 565, CZ-532 10 Pardubice, Czech Republic b Institute of Chemistry – Analytical Chemistry, Karl-Franzens University, Universitaetsplatz 1, A-8010 Graz, Austria Received 16 April 2007; received in revised form 18 July 2007; accepted 19 July 2007 Available online 31 July 2007

Abstract Hydrogen peroxide and nicotinamide adenine dinucleotide (NADH) may be determined amperometrically using screen-printed electrodes chemically modified with iron(III) hexacyanoosmate(II) (Osmium purple) in flow injection analysis (FIA). The determination is based on the exploitation of catalytic currents resulting from the oxidation/reduction of the modifier. The performance of the sensor was characterized and optimized by controlling several operational parameters (applied potential, pH and flow rate of the phosphate buffer). Comparison has been made with analogous complexes of ruthenium (Ruthenium purple) and iron (Prussian blue). Taking into account the sensitivity and stability of corresponding sensors, the best results were obtained with the use of Osmium purple. The sensor exhibited a linear increase of the amperometric signal with the concentration of hydrogen peroxide in the range of 0.1–100 mg L−1 with a detection limit (evaluated as 3σ) of 0.024 mg L−1 with a R.S.D. 1.5% for 10 mg L−1 H2 O2 under optimized flow rate of 0.4 mL min−1 in 0.1 M phosphate buffer carrier (pH 6) and a working potential of +0.15 V versus Ag/AgCl. Afterwards, a biological recognition element – either glucose oxidase or ethanol dehydrogenase – was incorporated to achieve a sensor facilitating the determination of glucose or ethanol, respectively. The glucose sensor gave linearity between current and concentration in the range from 1 to 250 mg L−1 with a R.S.D. 2.4% for 100 mg L−1 glucose, detection limit 0.02 mg L−1 (3σ) and retained its original activity after 3 weeks when stored at 6 ◦ C. Optimal parameters in the determination of ethanol were selected as: applied potential +0.45 V versus Ag/AgCl, flow rate 0.2 mL min−1 in 0.1 M phosphate buffer carrier (pH 7). Different structural designs of the ethanol sensor were tested and linearity obtained was up to 1000 mg L−1 with a maximum R.S.D. of 5.1%. Applications in food analysis were also examined. © 2007 Elsevier B.V. All rights reserved. Keywords: Screen-printed electrodes; Biosensor; Osmium purple; Ruthenium purple; Prussian blue; Hydrogen peroxide; Glucose; Ethanol; Flow injection analysis

1. Introduction Continuous and selective monitoring of various biological compounds such as glucose or ethanol is required in many fields of industry, especially in food processing. Common methods based on classic titration are time consuming; chromatographic techniques introduced recently require expensive laboratory equipment. In order to eliminate these drawbacks, the use of biosensors represents a low price alternative. These amperometric biosensors are based on enzymes that consume oxygen (all of the oxidases), produce hydrogen peroxide

∗

Corresponding author. Tel.: +420 466 037 512; fax: +420 466 037 068. E-mail address: [email protected] (K. Vytˇras).

0003-2670/$ – see front matter © 2007 Elsevier B.V. All rights reserved. doi:10.1016/j.aca.2007.07.053

(excluding oxidases producing water) or the reduced form of ␤-nicotinamide adenine dinucleotide NADH (dehydrogenases) during the course of the catalytic reaction on the substrate of interest [1]. One of the fundamental problems is the high overpotential necessary for the oxidation of H2 O2 (around +0.65 V) and NADH (+1.0 V) on a bare carbon electrode. Moreover, at the potential where the anodic oxidation takes place, various organic compounds, e.g., both ascorbic and uric acids, are co-oxidized. One of the possibilities how to overcome above problems is the use of mediators. Most common mediators used are ferrocene, ferricyanide, organic mediators like phenoxazines (Nile Blue, Meldola Blue), phenazines (phenazine methosulfate), phenathiazines (Toluidine Blue, Methylene Blue), quinones [2–4], numerous metal complexes, such as iridium [5], rhodium

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[6], ruthenium or osmium-complexes [7–10], Prussian blue and other metalhexacyanoferrates [11,12]. In our laboratory, we have demonstrated the suitability of such modification of SPEs with metal oxides, e.g., manganese dioxide [13], ruthenium dioxide [14], rhodium dioxide [15] and other oxides of the platinum metal group [16]. Iron hexacyanoosmate-modified electrodes based on carbon paste have already been studied for measuring the peroxodisulphates in hair-blonding boosters [17]. In addition, these types of complexes were first mentioned in connection with their structural studies [18] and investigations on the electrical properties [19]. To the authors’ knowledge, no report has been published on the application of iron hexacyanoosmate (Osmium purple) as a redox mediator or catalyst in connection with biosensors and therefore the modification of screen-printed carbon electrodes (SPCEs) with this complex for the determination of H2 O2 and NADH has been performed for the first time. In the present paper, the optimization is described to allow the determination of the above-mentioned analytes. Subsequently, the preparation of the biosensors for glucose or ethanol involving immobilisation of an appropriate enzyme is presented. From the oxidases, glucose oxidase (GOx) was used herein as a “model enzyme” referring to the fact that it is relatively inexpensive, easily available, and stable; from dehydrogenases we used ethanol dehydrogenase (ADH). A structural design of the ethanol sensor is also discussed and results of analyses of real samples are given. 2. Experimental 2.1. Materials Glucose oxidase (EC 1.1.3.4. from Aspergillus niger, specific activity 210 U mg−1 ; GOx), alcohol dehydrogenase (EC 1.1.1.1 from baker yeast, specific activity 316 U mg−1 ; ADH), Nafion (5% m/m solution in lower aliphatic alcohols), NAD+ and NADH were obtained from Aldrich. Chemicals used for the preparation of buffer, stock and standard solutions were of analytical reagent grade and purchased from Lachema (Brno, Czech Republic). All other reagents were of analytical grade (Merck). Phosphate buffer was prepared by mixing aqueous solutions of sodium dihydrogenphosphate and disodium hydrogenphosphate (both 0.1 M) to achieve solutions of the required pH. A glucose stock solution (2.5 g L−1 ) was prepared and diluted appropriately. Solutions of ascorbic acid and uric acid (both Aldrich, 50 mg L−1 ) were prepared immediately before use. For the evaporation of the solvent of the sample (for analysis with HPLC), nitrogen gas (purity 5.0, Linde Technoplyn a.s., Prague, Czech Republic) was used. 2.2. Instrumentation A modular electrochemical system, AUTOLAB, equipped with modules PGSTAT 30 and ECD (Ecochemie, Utrecht, Holland) was used in combination with a corresponding software (GPES, Ecochemie) under Windows® . The flow injection system consisted of a peristaltic pump (Minipuls 3, Gilson SA., France), a sample injection valve

(ECOM, Ventil C, Czech Republic), and a self-constructed thinlayer electrochemical flow-through cell. The working electrode was fixed via rubber gaskets (thickness 0.6 mm) directly to the back plate (counter electrode made of stainless steel) of the thin-layer cell with a Teflon support as a holder. The reference electrode was Ag/AgCl/3 M KCl (RE-6, BAS, USA), the stainless steel back plate represented the counter electrode of the cell. The responses were evaluated via peak heights (differences between background and response current of the analyte). For pH measurements a portable pH-meter (CPH 52 model, Elteca, Turnov, Czech Republic) equipped with a combined glass pH-sensor (OP-0808P, Radelkis, Budapest, Hungary) was used. The measuring cell was calibrated with standard buffer solutions of the conventional activity scale [20]. The glucose in honey was separated and quantified on an HPLC chromatograph equipped with a pump (SP 8800, SpectraPhysic, San Jose, USA), a degasser (Shodex Pegas KT-35M, Showa Denko KK, Tokyo, Japan), and an evaporative light scattering detector (ELSD; Chromachem® 6100, ESA Inc., Chelmsford, USA). 2.3. Procedure 2.3.1. Preparation of modifiers Osmium purple, Fe4 [Os(CN)6 ]3 (OP) was prepared from potassium osmate dihydrate (10 g), which was dissolved in water (10 mL) and mixed with a solution of potassium cyanide (10 g) in water (30 mL) and sodium hydroxide (30%, 0.1 mL). The mixture was then heated to boiling for half an hour after which another portion of 10 g KCN in 30 mL water was added. The temperature was kept at boiling for another 4–5 h. The resulting yellow solution was acidified with HCl to pH 2 (caution: all preparative steps involving KCN were performed in a well ventilated hood, due to the evolution of dicyanide and hydrogen cyanide). The product precipitated by addition of iron trichloride (3 g in 20 mL 0.01 M HCl) was isolated by centrifugation (10 min, 10000 rpm) and washed several times with water. The product was dried and stored in a dry place [17]. Ruthenium purple, Fe4 [Ru(CN)6 ]3 (RP). The precipitation was completely analogous to that of Osmium purple, using ruthenium trichloride hydrate (1.5 g) instead of the corresponding osmium compound. Prussian blue, Fe4 [Fe(CN)6 ]3 (PB). Potassium hexacyanoferrate(II) was dissolved in water and the solution acidified with HCl to pH 2. Iron trichloride dissolved in HCl (pH 2) was added while the mixture was stirred. The precipitate was isolated in the same way as the above complexes. 2.3.2. Electrode preparation Carbon ink (0.95 g, Gwent C2050617D12, Pontypool, UK) and corresponding hexacyanocomplex modifier (0.05 g) were thoroughly mixed manually for 5 min and subsequently sonicated for 30 min. The resulting mixture was immediately used for the fabrication of electrodes. The working electrodes were prepared by screen-printing modified inks onto an inert ceramic support (40 mm × 10 mm × 0.635 mm, ADS96R, CoorsTek, Glenrothes, UK). Thick layers of the modified carbon

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ink were formed by brushing the ink through an etched stencil (thickness 100 ␮m, electrode printing area 105 mm2 ) with the aid of the squeegee of the screen-printing device (Tesla UL 1505 A, Tesla Lanˇskroun, Czech Republic) onto the ceramic substrates. The resulting plates were dried at 60 ◦ C for ca. 2 h. For the GOx immobilization, 1 mg of the enzyme was dissolved in 20 ␮L of 0.1 M phosphate buffer (pH 7.5) and mixed with an equal amount of Nafion solution neutralized to pH∼7 with ammonia. The resulting mixture (5 ␮L) was applied directly onto the active area of the OP bulk-modified SPCE surface and air-dried. The ethanol sensor containing alcohol dehydrogenase (ADH) and nicotinamide adenine dinucleotide (NAD+ ) was constructed via two different procedures: T1: 50 ␮L of a mixture of 2 mg ADH and 10 mg NAD+ dissolved in 10 mL phosphate buffer (pH 7.0) was added to 5 mL of the sample and directly injected into the carrier stream; T2: 5 ␮L of a mixture (2 mg NAD+ dissolved in 100 ␮L phosphate buffer, pH 7.0) was dropped onto the active electrode surface. For enzyme immobilization, 1 mg of the enzyme (ADH) was dissolved in 20 ␮L of 0.1 M phosphate buffer (pH 7.0) and mixed with an equal amount of 0.05% Nafion solution neutralized to pH∼7 with ammonia. The resulting mixture (5 ␮L) was applied directly onto the layer of NAD+ and left to dry. Prior to measurements, the modified screen-printed carbon electrodes were pre-treated by scanning the potential several times between −0.8 and +1.0 V in a 0.1 mol L−1 phosphate buffer (pH 6.5) with the applied scan rate of 100 mV s−1 . 2.3.3. Optimization process The measurements of H2 O2 , NADH, glucose and ethanol were performed by employing DC amperometry. At the outset, it was important to optimize all operational variables, e.g. applied potential (from +0.6 to −0.3 V versus Ag/AgCl), pH of phosphate buffer (5–9) and flow rate (0.1–1.0 mL min−1 ). 2.3.4. Analysis of real samples Flow injection measurements of glucose were carried out in 0.1 M phosphate buffer (pH 6.0). The applied potential was set at +0.15 V versus Ag/AgCl with a typical flow rate of 0.2 mL min−1 , and the injected sample volume was 200 ␮L. In case of the ethanol sensor, the T1 type arrangement was used to study the concentration of ethanol in beverages under optimized conditions: applied potential +0.45 V versus Ag/AgCl, flow rate of phosphate buffer (pH 7.0) 0.2 mL min−1 . Glucose was separated on a Purospher Star® NH2 column (4 mm × 250 mm, Merck, Darmstadt, Germany) at 35 ◦ C with a column heater (LCO 101, ECOM, Prague, Czech Republic). A flow rate 1.0 mL min−1 was used with the mobile phase acetonitrile/water (80:20, v/v). The Chromachem® 6100 detector was set at 45 ◦ C (evaporation temp.) and 30 ◦ C (nebulisation temp.) with 1.8 × 105 Pa of nitrogen gas. The mobile phase was filtered through a membrane filter (0.45 ␮m, 47 mm, Merck, Darmstadt, Germany) and degassed in an ultrasonic bath (SONOREX RK 52 H, Bandelin electronic GmbH & Co. KG, Berlin, Germany). The injection volume of sample was 10 ␮L. The appropriate retention time was 6.8 min for glucose.

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2.3.5. Sample processing Food and drink samples were bought in a local shop in Pardubice (CZ) and stored at room temperature (honey, vodka, plum brandy). Samples were appropriately diluted with phosphate buffer and injected directly into the buffer carrier stream for analysis. 3. Results and discussion 3.1. Sensors based on oxidases 3.1.1. Response to hydrogen peroxide Hydrogen peroxide is a by product of the biocatalytic reactions involved with a majority of oxidases. Therefore, attention was paid first to optimize the conditions of the H2 O2 determination. To cover all operational and experimental variables affecting the amperometric determination of hydrogen peroxide, the dependence of the signal on the working potential, the pH of supporting electrolyte, the flow rate, etc. was studied in detail. Electrochemical oxidation of H2 O2 was investigated at both unmodified SPCEs and those modified with OP, PB and RP. No or very low response of hydrogen peroxide was observed at unmodified SPCE in the whole examined range (−0.3 to +0.6 V) (not shown). The behaviour of H2 O2 at SPCEs modified with OP reveals that it produces a catalytic current response mainly due to the reduction (Fig. 1). Thus, the reduced form of the modifier is oxidized by hydrogen peroxide yielding more Fe4 III [Os(CN)6 ]3 at the surface of the electrode than is physically present. In hydrodynamic voltammograms, a well developed cathodic peak for H2 O2 with maximum at +0.15 V can be observed. As expected, the effect is similarly established for the other complex, PB, reaching its maximum catalytical activity at −0.15 V as almost all the ferro-/ferricyanide moieties of PB are known to be electroactive with fast kinetics. However, RP yielded notably lower sensitivity and proved not to be suitable for such studies. Therefore, SPCEs modified with OP are appropriate for the determination of H2 O2 and, as a consequence, the following studies were performed with this complex only.

Fig. 1. Dependences of the FIA peak current of H2 O2 on the applied potential for different mediators. Carrier phosphate buffer pH 6.5 (0.1 mol L−1 ); flow rate 0.4 mL min−1 ; injection volume 200 ␮L of H2 O2 (10 mg L−1 ).

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To optimize the analytical conditions, the effect of the pHvalue on the signal response was studied (not shown). Within the range of pH 5 and 7, the response was the highest and remained constant. At higher pH values, the responses significantly decreased. This could be attributed to the dissolution of mediator and its leaching out of the electrode. The flow rate has only a minor influence on the height of the response; it showed a maximum and constant value of the signal response within 0.4–0.8 mL min−1 . With respect to analysis time and sample throat, a flow rate of 0.4 mL min−1 was chosen as an optimum. Under such conditions, a linear relation between the current and concentration of hydrogen peroxide was found in the range from 0.1 to 100.0 mg L−1 , which is in a good correlation with sensors modified with nickel hexacyanoferrate [12]. The R.S.D. was determined as 1.5% for a concentration of 10 mg L−1 hydrogen peroxide (n = 5) with a detection limit of 24 ␮g L−1 . For comparison, Fig. 2 shows calibration curves of SPCEs modified with all complexes. 3.1.2. Determination of glucose Glucose oxidase was chosen as a model enzyme in this study considering the fact that it is relatively inexpensive, readily available and robust. Additionally, it belongs to the most utilized enzymes in bioscience research and development, food industry and especially in biomedicine for monitoring glucose in blood of patients suffering from diabetes mellitus. SPCEs modified with OP were additionally modified with GOx entrapped in Nafion. The experimental conditions were optimized again for the biosensor, because the presence of enzyme and the membrane could change the electrochemical properties of the sensor. The optimum operational potential was influenced minimally by the presence of the biocatalyst. However, with negative potentials (below 0.0 V) surprisingly the oxidation current increased again (Fig. 3). We assume two different mediated/catalytic mechanisms. The first part from +0.6 V to +0.0 V is based on oxidation/reduction of H2 O2 with a response time of around 30 s. The second part (from +0.0 V to −0.3 V) had slower responses, 40 s or more. Probably the reason for this are two systems of com-

Fig. 2. Calibration graph for hydrogen peroxide. Applied potential +0.15 V (OP), −0.1 V (PB), −0.2 V (RP) vs. Ag/AgCl; carrier 0.1 M phosphate buffer pH 6.0 (OP, RP), 8.0 (PB); flow rate 0.4 mL min−1 ; injection volume 200 ␮L.

Fig. 3. Dependences of the FIA peak current of glucose on the applied potential. Carrier phosphate buffer pH 6.0 (0.1 mol L−1 ); flow rate 0.4 mL min−1 ; injection volume 200 ␮L of glucose (50 mg L−1 ).

plexed osmium species, maybe Os(VI/IV) and Os(III/II). From both, the higher oxidation state may oxidize hydrogen peroxide to oxygen (producing oxidation currents at higher potentials), whereas the lower is oxidized by the analyte to the higher oxidation state (generating reduction currents at lower potentials). A further decrease of the potential shifts from one system to the other with superimposition of both effects leading to an intermediate decrease of the reduction current (or even occurrence of oxidation currents). Similar results were obtained with other mediators/catalysts containing platinum metal oxides [15,16]. For flow injection analysis (FIA), the modified electrode was used as an amperometric detector in a thin-layer cell with the supporting electrolyte as a carrier. In contrary to hydrogen peroxide, the highest response was achieved with 0.1 mL min−1 (for the range 0.1–1.0 mL min−1 ) and decreased exponentially up to 1.0 mL min−1 . Such behaviour refers to a dependence of the electrochemical reaction rate on the enzymatic reaction and the penetration through the membrane. For our purposes, the flow rate was set at 0.2 mL min−1 . With respect to the dependence of the current on pH, similar results were obtained with glucose and hydrogen peroxide, as expected. The selected experimental conditions (operation potential +0.15 V, flow rate 0.2 mL min−1 , pH 6) were used to establish the calibration curve, showing linearity between the current response and the concentration of glucose in the range from 1 to 250 mg L−1 . At higher concentrations, deviation from linearity was observed and the current remained constant beyond 500 mg L−1 (Fig. 4) probably due to a saturation of the active centres in the enzyme. The detection limit (3σ) was found to be 20 ␮g L−1 , the R.S.D. was determined as 2.4% (n = 5) for a concentration of 100 mg L−1 of glucose. The mechanical reproducibility among different sensors fabricated was around 5.6%. No loss of the original signal was achieved after 3 weeks, when stored at 6 ◦ C in the refrigerator. The glucose biosensor described above was applied to determine the glucose content in a sample of honey; the measurements were assessed using a calibration graph. As can be seen (Table 1), the results are in accordance with the reference HPLC method and the standard representation of the glucose content. The

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Fig. 4. Calibration graph for glucose. Applied potential +0.15 V vs. Ag/AgCl; carrier 0.1 M phosphate buffer pH 6.0; flow rate 0.2 mL min−1 ; injection volume 200 ␮L.

obtained results encourage the possible use of the proposed sensor for determining glucose in such types of food samples. 3.2. Sensors based on dehydrogenases More than 250 dehydrogenases are known which require nicotinamide coenzymes as co-reactants. Amperometric biosensors based on nicotinamide adenine dinucleotide (NAD+ /NADH) dependent dehydrogenases can be used to monitor a variety of biologically important molecules in biotechnology and analysis. Sensors based on the oxidation of the substrate with the participation of dehydrogenases create proportional amounts of detectable NADH from NAD+ [21] 3.2.1. Response to NADH Using the sensor modified with OP, NADH was found electroactive at anodic potentials only with the oxidation response starting at approx. +0.25 V (Fig. 5). At unmodified SPEs, well-defined peaks were observed with NADH, however, the responses to NADH considerably deteriorated with the number of injections, perhaps in consequence to passivation of the electrode by the adsorption of the reaction product. On the other hand, the signals of the OP-modified electrode were stable with a R.S.D. 1.5% with a concentration of 0.01 mM NADH (n = 3). The effect of pH upon the peak current for NADH is demonstrated in Fig. 6. It can be seen that the changes of pH between 5 and 7.5 have a relatively small effect. A substantial increase of the signal is observed at pH 8, and the response levels off with higher pH. However, at pH 8 responses to NADH were unstable

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Fig. 5. Dependences of the FIA peak current of NADH on the applied potential. Carrier phosphate buffer pH 7.45 (0.1 mol L−1 ); flow rate 0.2 mL min−1 ; injection volume 200 ␮L of NADH (1 mM).

and decreased with the number of injections caused by dissolution of the modifier as mentioned above. The highest response to NADH at higher pH can be explained by the fact that at the electrode surface due to the decomposition of the mediator there was partially dissolved OP and therefore better contact among electrode, complex and NADH was ensured. To avoid any loss of the stability of the sensor, a supporting electrolyte of pH 7 was chosen as an optimum. Employing the operational parameters discussed above (+0.45 V, flow rate 0.2 mL min−1 , pH 7.0), a linear relation between the concentration of NADH and the current response was found in the range 0.001–0.1 mM. At concentrations higher than 0.1 mM, only moderate deviation from linearity was observed (Fig. 7). 3.2.2. Ethanol Such a sensor responding to NADH may be exploited for the determination of quite a number of biologically active substances after immobilisations of corresponding dehydrogenases.

Table 1 Determination of glucose in honey Sample

Method

n

x (%)

R.S.D. (%)

Honey

Present HPLC Conventional content

3 3

32.5 35.5 26.0–33.0

2.2 1.4

Operational potential +0.15 V vs. Ag/AgCl; carrier phosphate buffer pH 6.0 (0.1 mol L−1 ); flow rate 0.2 mL min−1 ; injection volume 200 ␮L. x, arithmetic mean; n, number of replications; R.S.D., relative standard deviation.

Fig. 6. Effect of pH. Applied potential +0.45 V vs. Ag/AgCl; carrier phosphate buffer (0.1 mol L−1 ); flow rate 0.2 mL min−1 ; injection volume 200 ␮L. NADH concentration 0.1 mM.

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P. Kotzian et al. / Analytica Chimica Acta 599 (2007) 287–293 Table 2 Determination of ethanol in beverages Sample

n

x (%)

R.S.D. (%)

Declared (%)

Plum brandy (Slivovice) Vodka

3 3

48.8 43.2

6.8 6.8

52.0 42.0

Operational potential +0.45 V vs. Ag/AgCl; carrier phosphate buffer pH 7.0 (0.1 mol L−1 ); flow rate 0.2 mL min−1 ; injection volume 200 ␮L. x, arithmetic mean; n, number of replications; R.S.D., relative standard deviation.

Fig. 7. Calibration graph for NADH. Applied potential +0.45 V vs. Ag/AgCl; carrier 0.1 M phosphate buffer pH 7.0; flow rate 0.2 mL min−1 ; injection volume 200 ␮L.

Initial studies were performed with the enzyme ethanol dehydrogenase. In this work, the main focus was preferably put on the construction of the sensor because the dehydrogenase requires the presence of the soluble cofactor (NAD+ ) and therefore, problems with its leaching from the electrode in FIA may arise. First, we tried to add a mixture of the enzyme and cofactor (T1) to the ethanol sample. Reproducible responses were obtained (R.S.D. = 1.3%, c = 1000 mg L−1 , n = 3), however, the disadvantage was that both the enzyme and the cofactor had to be added separately for each sample which was time- and reagents-consuming. A linear relation between the concentration of ethanol and the current response was found in the range from 100 to 500 mg L−1 (Fig. 8). Although both the enzyme and the cofactor are expensive, the costs for one sample sum up to around 0.07 USD only. To shorten the analysis time (and costs for analysis as well), immobilisation of the enzyme together with NAD+ by the aid of Nafion was tested. A layer-type arrangement was found important because large molecules, such as FAD or NAD+

cannot pass freely through the Nafion membrane, or at least their diffusion is significantly restricted. If both, the enzyme ADH and NAD+ were immobilised together in Nafion, no response for ethanol was achieved. Therefore, the cofactor was first adsorbed directly on the surface of the SPE (T2) with a second layer formed by Nafion with the entrapped enzyme. Such an ethanol sensor showed a calibration curve with linearity between current and concentration of alcohol in the range of 100–1000 mg L−1 (Fig. 8). However, the reproducibility (R.S.D. = 5.1%, c = 1000 mg L−1 , n = 3) and the stability of such an arrangement were poor and recalibration for each sample was needed, probably caused due to degradation of the enzyme. The ethanol sensor (T1) was applied to the determination of the ethanol content in two types of beverages; measurements were assessed using a calibration graph. As can be seen, the results are acceptable and correspond with the declared values (Table 2). 4. Conclusion A sensor based on screen-printed electrodes modified with iron hexacyanoosmate exhibited good stability, enhanced electrocatalytic activity towards H2 O2 and NADH. Moreover, Osmium Purple had even slightly higher sensitivity towards hydrogen peroxide than commonly used Prussian blue. In case of the detection of NADH, one big advantages of the modifier is its practical insolubility in water. Therefore, FIA could be used without any protective membrane in contrary to the widely used Meldola Blue mediator, which significantly reduces the analysis time. Acknowledgements This work was supported from the Ministry of Education, Youth and Sport of the Czech Republic (projects MSM0021627502 and LC 06035) and from the Czech Science Foundation (No. 203/05/2106). References [1] [2] [3] [4] [5]

Fig. 8. Calibration graph for ethanol. Applied potential +0.45 V vs. Ag/AgCl; carrier 0.1 M phosphate buffer pH 7.0; flow rate 0.2 mL min−1 ; injection volume 200 ␮L. Other conditions as in the text.

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