Kinetics of oxidation of hydrogen peroxide at hemin-modified electrodes in nonaqueous solvents

Bioelectrochemistry 76 (2009) 63–69

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Bioelectrochemistry j o u r n a l h o m e p a g e : w w w. e l s ev i e r. c o m / l o c a t e / b i o e l e c h e m

Kinetics of oxidation of hydrogen peroxide at hemin-modified electrodes in nonaqueous solvents☆ Zuzana Brusova, Edmond Magner ⁎ Department of Chemical and Environmental Sciences, Materials and Surface Science Institute, University of Limerick, Limerick, Ireland

a r t i c l e

i n f o

Article history: Received 18 December 2008 Received in revised form 11 February 2009 Accepted 27 February 2009 Available online 13 March 2009 Keywords: Hemin Carbon electrodes Nonaqueous solvents Peroxide

a b s t r a c t Hemin adsorbed on graphite electrodes and used to catalyse the reduction of hydrogen peroxide in an aqueous buffer and in a range of nonaqueous solvents has been described. The immobilised hemin is stable in the solvents examined. The rate limiting step involves the reaction between hemin and hydrogen peroxide. Kinetic analysis of the response in nonaqueous solvents showed that Imax / Kapp m increased linearly with the solvent hydrophobicity (log P) in all solvents, a trend that is explained by preferable partitioning of hydrogen peroxide into the polar hemin layer. © 2009 Elsevier B.V. All rights reserved.

1. Introduction Hydrogen peroxide and peroxides in general are important reagents in many biological, environmental and industrial processes [1]. Hydrogen peroxide is a major oxygen reactive species in living organisms. Oxidative damage resulting from the cellular imbalance of H2O2 and other reactive oxygen species is connected with aging and severe human diseases such as cancers and cardiovascular disorders [2]. Organic hydroperoxides formed by oxidation of lipids by free radicals can be found in vegetable oils, baby foods and biological tissue [3]. Peroxides present in living cells can lead to cell damage and resulting development of cancer as mentioned above. Hydroperoxides present in food accelerate nutrition damage, generate unpleasant odours and may have toxic effects [4], e.g. rancidification of fats [5]. Various techniques are available for the determination of peroxide content in samples, e.g. iodometric, coulometric and amperometric detection, chemi-luminiscence, polarographic and spectroscopic methods [6,7]. However, these methods can have a number of disadvantages, such as low sensitivity, interference effects and a requirement for complicated and/or time consuming sample pre-treatment. In the last two or three decades significant effort has been devoted to the development of biosensors for the detection of hydrogen peroxide [7–12] and organic hydroperoxides [3,4,6,13–17]. Some of these biosensors have been successfully applied to the characterisation of samples such as creams, lotions, olive oils etc. [4,12,17].

☆ Dedicated to Professor Lo Gorton on the occasion of his 60th birthday. ⁎ Corresponding author. Tel.: +353 61 202628; fax: +353 61 213529. E-mail address: [email protected] (E. Magner). 1567-5394/$ – see front matter © 2009 Elsevier B.V. All rights reserved. doi:10.1016/j.bioelechem.2009.02.014

Peroxidases (horseradish peroxidase, HRP, chloroperoxidase, soybean peroxidase [12], palm tree peroxidase [18]) have been the enzymes of choice for detection of peroxides. The need to assay hydrophobic substrates (organic hydroperoxides, cholesterol, pesticides etc.) which are insoluble in aqueous media, together with the finding that enzymes retain their catalytic activity in organic media [19,20], led to the development of organic phase enzyme electrodes (OPEEs) [14,15,21]. In addition to the ability to analyse hydrophobic samples, OPEEs have often enhanced thermostability, sensitivity and linear ranges when compared to biosensors working in aqueous media. Microbial contamination is eliminated and unwanted side reactions (e.g. hydrolytic reactions) are suppressed. Despite the numerous advantages of using enzyme biosensors in organic media and enzyme biosensors in general, certain drawbacks result directly from the enzyme properties. Enzymes are expensive and relatively unstable due to deactivation. In organic media, the majority of enzymes possess only a fraction of the catalytic activity observed in water [22]. In many cases transfer of the enzyme into organic media can lead to inactivation denaturation [19]. The majority of enzymes used in biosensors are bulky molecules with redox active centres deeply buried within an insulating protein matrix and hence inaccessible resulting in slow rate of reason direct electron transfer (DET). A range of methods to direct the orientation of the enzyme on the electrode surface have been suggested but satisfactory results are often difficult to achieve and mediators are widely used in order to establish reasonable rates of electron transfer. Finally, immobilisation of enzymes onto the electrode surface during the construction of a biosensor may lead to perturbation in the microenvironment of the enzyme active site or significant conformational changes in the enzyme structure causing partial or full enzyme denaturation, e.g.

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during adsorption, denaturation is likely due to multiple contacts and interactions of the enzyme with the electrode surface. Hemin (iron protoporphyrin IX) is the active centre of many haemcontaining redox proteins. Due to its small size, direct electron transfer can be established independently of the orientation on the electrode surface [23] while higher surface coverages should lead to improved sensitivity (current density) and response when hemin is used in a sensor [10]. Portsmouth et al. proposed that the mechanism for the peroxidase activity of deuterohemin was basically the same as the accepted peroxidase mechanism, but with some differences [24]. Hemin did not undergo degradation, even at high peroxide concentrations in the presence of a second substrate (hydrogen donors), showing that the catalyst was regenerated during the reaction. The primary active haem–peroxide complex (compound I) was formed at a much faster rate in peroxidases (ca. 3000-fold) than in hemin. Also, no intermediate analogous to peroxidase compound II was observed in the case of deuterohemin. In the reaction of the hemin–peroxide complex with a second substrate, only one (possibly a 2-electron) step could be distinguished, whereas in peroxidases it is generally known to proceed via two 1- electron reactions. The bioelectrochemical reduction of H2O2 was observed by Lötzbeyer et al. with HRP-, myoglobin- and cytochrome c-modified electrodes, as well as with hemin- and MP-11-modified electrodes [10,25]. Smaller molecules, where the redox centre is more readily accessible to the substrate were more efficient electrocatalysts. The highest electrocatalytic efficiency for H2O2 reduction was observed with hemin, higher by a factor of 30 when compared to MP-11 [10] and by a factor of about 18,000 when compared to immobilised HRP [25]. This is in contrast to homogeneous enzymatic catalysis, where HRP was about 3000 fold more active than hemin [10,24] or other smaller molecules, and can be explained by the fact that the large size of the HRP molecule was optimized by nature [10]. Sensors based on direct electron transfer bring numerous advantages when compared to sensors using mediators. Leakage of the mediator from the sensor does not have to be considered. Also, possible limitations of the overall reaction rate by diffusion-limited mass transport of the mediator can be ruled out [10]. The concept of minizymes, and mainly hemin alone, can be advantageous when compared to using enzymes such as HRP for sensor development. Faster rates of DET are observed, hemin is considerably cheaper than enzyme preparations and the possibility of protein denaturation by the organic solvent does not arise. A number of reports have described the use of hemin as a catalyst in organic media. A PEG-modified hemin complex was prepared [26] with a ca. 100-fold higher catalytic activity in organic solvents that in aqueous buffer. The PEG–hemin preparation was soluble both in water and organic solvents, and showed a spectrum with a sharp Soret band at 398 nm in trichloroethane, typical for hemin solutions. The complex was soluble in water as well as in organic solvents such as benzene. It displayed peroxidative catalytic activity in anhydrous benzene for the oxidation of o-phenylenediamine using hydrogen peroxide or t-butyl hydroperoxide as oxidant. The haem–surfactant complex showed significantly enhanced stability towards hydrogen peroxide when compared to unmodified hemin. This increased stability was ascribed to the surfactant stabilising the active centre in a manner analogous to that of the apoprotein in the native enzyme [27]. The insolubility of hemin in neutral aqueous media limits its applications as an effective homogeneous catalyst. However, using hemin for homogeneous nonaqueous catalysis broadens the range of possible applications. An alternative method, which overcomes the insolubility of hemin in water and many organic solvents, is to immobilise hemin onto a suitable support. In this report, we describe the properties of hemin immobilised on graphite electrodes and examine the catalytic activity of the immobilised hemin in a range of nonaqueous solvents.

2. Experimental 2.1. Materials Hemin chloride, potassium hydrogen phosphate, potassium dihydrogen phosphate, potassium chloride, Trizma® Base, ethyl acetate (99.8% HPLC grade), 1-pentanol (99%), toluene (anhydrous, 99.8%), tetraethylammonium-p-toluensulfonate (TEATS), lithium perchlorate, tetrabutylammonium tetrafluoroborate (TBATFB), and trifluoroacetic acid (TFA) (99%), acetone (99.8% Chromatosolv®) and 1hexanol (anhydrous, 99+%) were from Sigma-Aldrich. Acetonitrile (N99.8%), 1-octanol (99.5%), ethanol (absolute) and tetraoctylammonium bromide (TOABr) were purchased from Fluka. Methanol (99.9%), 1-propanol (N99%), 1-butanol (N99.7%), N,N-dimethylformamide (DMF, 99.9% Spectranal®), tetrahydrofuran (THF, 99.9% Chromatosolv®) and hydrogen peroxide (30%) were from Riedel de Haën. Nitric acid (69%), agar (fine powder), hydrochloric acid (35–37%) and sodium carbonate were purchased from BDH, England. Alumina for electrode cleaning was supplied by CH Instruments (0.05 µm Gamma and 1.0 µm Alpha). All aqueous solutions were prepared from deionised water (N18.2 MΩ) from an Elgastat Purification system. Organic solvents were used as received. Organic solvent–aqueous mixtures were prepared by mixing 95:5 % v/v ratio of the organic solvent containing 0.1 M electrolyte and 0.1 M TRIS buffer, pH 7.0. All solutions for cyclic voltammetry experiments were purged with nitrogen for 60 min. Fresh hydrogen peroxide stock solutions (0.1 M and 1.0 M) were prepared daily in the corresponding solvent. 2.2. Electrode preparation Prior to immobilisation of hemin, glassy carbon electrode tips were polished using alumina (1 μm and 0.05 μm particle size) and rinsed extensively with water, acetone and water. Occasionally, electrodes were dipped into concentrated nitric acid prior to polishing to achieve complete removal of strongly adsorbed hemin. Any residual material was removed from the electrode surface by gentle polishing on soft Kimcare Medical Wipes (Kimberly-Clark®) and rinsed with deionised water. 5 μl of 0.5 mg/ml hemin solution in 0.01 M carbonate buffer pH 10 was then placed onto the dried electrode surface and the electrodes were left to dry for at least 3 h. The hemin solutions were prepared fresh daily. Prior to voltammetry experiments, electrodes were rotated in carbonate buffer pH 10 (10 min, 2000 rpm) in order to remove loosely bound hemin. The measured current is reported as current density per surface concentration of hemin (A mol). 2.3. Electrochemical measurements Electrochemical experiments (cyclic voltammetry and amperometry) were performed with a CH Instruments (USA) potentiostat CHI630A with corresponding CHI630 software. 95–100% iR drop compensation was employed in all voltammetric experiments. A large volume (50 ml) closed electrochemical cell, kept under nitrogen atmosphere was used for voltammetric experiments. For amperometric experiments a glass 30 ml electrochemical cell was used. In amperometric experiments the reference electrode was separated in a second compartment and connected via salt bridge (a glass tube connected to Tygon® tubing, filled with the agar gel, and sealed by a Vycor® tip on the glass side, which was immersed in the working electrode compartment containing organic solvents and hydrogen peroxide). The counter electrode was also placed in the reference compartment for easier manipulation. A glassy carbon rotating disc electrode (3 mm diameter) mounted onto a rotating disc electrode system (Metrohm 628-10, Herisan, Switzerland) was used as the working electrode. Ag|AgCl (sat. KCl) electrode and a platinum wire were used as the reference and counter

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electrodes, respectively. All potentials are reported with respect to Ag| AgCl. Experiments were carried out at room temperature of 18 ± 1 °C. Amperometric detection of hydrogen peroxide hemin-modified electrodes was performed at an applied potential of +100 mV vs. Ag|AgCl electrode and a rotation rate of 2000 rpm. The first aliquot of hydrogen peroxide was added after 600 s to enable stabilisation of the baseline current. A new electrode was used for each experiment unless otherwise stated, and at least three electrodes were used for each measurement.

2.4. HPLC measurements HPLC was performed on a Waters system equipped with Breeze software (version 3.20). Isocratic elution was used with a flow rate of 1 ml/min. The samples (10 or 20 µl sampling volume) were injected using a Waters 717+ autosampler. A Waters 1525 binary pump was used with a reverse phase column (Waters Xterra C18, 4.6 × 100 mm). Hemin was detected using a Waters 2487 Dual λ detector at 393 nm. The mobile phase consisted of 0.1 M TFA in water mixed with 0.1 M TFA in acetonitrile in 60:40 v/v ratio and was vacuum filtered before use. UV–vis spectroscopy experiments were performed with a Shimadzu 1201 spectrophotometer.

2.5. Stability of hemin-modified electrodes Hemin-modified glassy carbon electrodes were prepared as described previously. When dried, they were rinsed with deionised water and exposed to a series of stability tests (2000 rpm, open circuit potential) in various solutions: (1) 10 min rotation in carbonate buffer pH 10, (2) 20 min rotation in phosphate buffer pH 7, (3) when testing in phosphate buffer, rotation for 20 min (×3), (4) when testing in 95% ethanol (containing 5% TRIS buffer pH 7) solutions, rotation for 20 min (×2 in 95% ethanol and 20 min in phosphate buffer, (5) when testing in hydrogen peroxide, rotation for 15 min (×2) in 50 µM H2O2 solution and 15-minute rotation in 150 µM H2O2, (6) in 95% ethanol containing hydrogen peroxide, the same procedure as in (5) was used with the phosphate buffer solution being replaced by 95% ethanol. After each of the five rotation cycles the electrode was rinsed with deionised water, allowed to air dry after which the surface coverage was determined by cyclic voltammetry in deaerated phosphate buffer (K-PBS, pH 7).

Fig. 2. HPLC chromatograms of (A) freshly prepared, (B) 1 day old and (C) 2 week old hemin solutions.

2.6. Determination of water content The amount (w/w) of water present in organic solutions was determined using a Karl Fischer titrator, Mettler Toledo DL31. 3. Results and discussion 3.1. Hemin stability

Fig. 1. Visible spectra of hemin solution in aqueous buffer pH10: fresh ( ), 1 day ( ), 2 days ( ), 5 days, ( ), 2 weeks old ( ), and fresh hemin dissolved in the mobile phase for HPLC ( ), see the experimental section. Inset: 450–700 nm region.

The stability of hemin in solution was examined by UV–vis spectroscopy (Fig. 1). When hemin was dissolved in the HPLC mobile phase, the spectrum displayed a sharp Soret band (399 nm), Q bands (501 and 539 nm) and a charge transfer band (627 nm), and was very similar to the spectrum reported by Laszlo et al. [28]. Upon dissolution of hemin in carbonate buffer pH 10, the Soret band intensity decreased, broadened and moved to ca. 390 nm. These changes together with the appearance of a shoulder at 360–370 nm are typical of the µ-oxo dimers formed in aqueous solutions [27]. The shift in the CT-band wavelength to 613 nm for the fresh hemin solution, and to 608–600 nm for the older solutions, may be explained by the presence of ferric porphyrin complex axially ligated by an anionic oxygen ligand [28], further confirming the presence of µ-oxo dimers. However, the spectra for fresh hemin solutions (both in carbonate buffer and the mobile phase, Fig. 1 inset), have a high degree of similarity, but differ significantly from the spectra of older solutions, indicating that changes in the hemin solutions are due to aging of the solution and not the solvent used. The intensity of the Soret band further decreased and the shoulder at 365 nm became more pronounced with time,

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indicating further hemin aggregation. The disappearance of the Q bands from the spectra of solutions which were over 1 day old also indicated that significant changes to the structure had occurred in solution. The stability of the hemin solutions was examined by HPLC. A freshly prepared solution of hemin was compared with solutions that were prepared earlier (1 day and 2 weeks). Significant changes were observed when comparing the fresh and older solutions. The relative area of the hemin peak obtained by HPLC decreased from 1.1 × 107 for the fresh hemin solution to 3.4 × 105 for the 1 day old sample, and to 1.5 × 105 for the 2 week old sample. Under the experimental conditions used, hemin elutes at 8.6 min as a single peak. In the fresh solution only monomeric hemin was present, (Fig. 2A). With time, the amount of monomeric hemin decreases and other products of hemin degradation appeared, (Fig. 2B and C). After 2 weeks, the amount of early eluting impurities increased significantly and the amount of monomeric hemin was negligible (Fig. 2C). If only dimerisation of hemin occurred, the dimers would be most probably dissolved by the organic solvent present in the mobile phase and a single peak would be observed even for older solutions. However, other products of hemin degradation are present even after 1 day, indicative of low stability of aqueous hemin solutions. Therefore, hemin solutions were prepared freshly each time immediately prior to electrode modification.

3.2. Surface coverage and operational stability of hemin electrodes Hemin adsorbs strongly onto carbon electrodes and in particular onto pyrolytic graphite [29,30]. The preparation of a hemin-modified electrode is simple and fast, and no extensive electrode polishing or other requirements for oriented immobilisation are necessary. Cyclic voltammograms indicate that the response is quasi-reversible with peak separations of 50 mV (Fig. 3). Plots of peak current versus scan rate were linear (data not shown), indicative of a surface confined species. The surface coverage of hemin was determined from integration of the peak current, yielding a value of 135 ± 14 pmol/ cm2 (n = 50 electrodes) which is in good agreement with previous reports of 160 pmol/cm2 [30] and 150 pmol/cm2 [31]. The area of a molecule of hemin is approximately 2.4 nm2, yielding a value of 75 pmol cm− 2 for monolayer coverage [32]. The measured value of

Fig. 4. Plot of response (current density per surface concentration of hemin) versus concentration of hydrogen peroxide in 90% ( ), 95% ( ), 98% ( ) and 99% v/v ( ) ethanol solutions.

135 pmol cm− 2 corresponds to slightly less than monolayer coverage, given the surface roughness of the graphite electrodes [33]. In phosphate buffer, the amount of hemin on the glassy carbon surface gradually decreased and reached on average 87% of the original value after five rotation cycles (1 cycle consisted of 20 min rotation in phosphate buffer, 2000 rpm, open circuit potential, surface coverage measured in deaerated buffer). The same trend was observed for electrodes rotated in hydrogen peroxide with the final surface coverage decreasing by 15% after 5 cycles. In the presence of both ethanol and hydrogen peroxide the decrease in the response due to hemin loss and/or degradation was more pronounced, about 8% after each cycle. However such stability is sufficiently high to test the electrodes in organic solvents containing hydrogen peroxide, if the duration of the experiment after peroxide addition is minimised to several minutes and a new electrode is used in each experiment. 3.3. Water content It is generally established that enzymes require some essential water for activity in organic solvents [20]. The water layer around the enzyme works as a lubricant, giving the enzyme the necessary flexibility to be catalytically active. Enzymes functioning in polar organic solvents generally require more water (typically around 2–5%), as the organic solvent can strip off the surface and active site bound water molecules, penetrate into the active site and cause denaturation of the enzyme. In contrast, the lack of water stripping ability in hydrophobic solvents allows efficient hydration of the enzyme active site at much lower amounts of added water (less than 1%). Even though it is well established that the catalytic cycle of peroxidases (and possibly hemin) is more complex than the Michaelis–Menten mechanism [34,35], it is useful to utilise this simplified approach to evaluate the apparent values for comparative purposes. The rate of reaction can be expressed in terms of the catalytic current [35]: app


Fig. 3. Cyclic voltammograms of hemin-modified glassy carbon electrode at a hemin surface coverage of 135 pmol/cm2. Scan rates 5, 2, 1 and 0.5 V/s outer to inner voltammograms.

Ik = Imax ½H2 O2  = ½H2 O2  + Km

ð1Þ

Imax = nFAk2 C

ð2Þ

is the apparent Michaelis constant for adsorbed hemin where Kapp m (Kapp m and [H2O2] in units of mM), Ik is the kinetic current, Imax is the maximum catalytic current (Ik and Imax in units of Amol), F is the Faraday (in unit of A s mol− 1), A is the surface area of the electrode (cm2), k2 is the rate constant for the conversion of enzyme–substrate

Z. Brusova, E. Magner / Bioelectrochemistry 76 (2009) 63–69 Table 1 Kinetic characteristics for H2O2 reduction by hemin electrodes in aqueous ethanol solutions. EtOH (% v/v)

Sensitivity (A mol− 1/mM)

−Imax (A mol− 1)

Kapp m (mM)

Imax / Kapp m (A mol− 1/mM)

90 95 98 99

1710 ± 120 1380 ± 90 890 ± 90 540 ± 110

27690 ± 3780 11510 ± 190 6190 ± 550 2930 ± 460

15.1 ± 1.3 7.5 ± 0.6 6.4 ± 0.4 4.7 ± 0.2

1830 ± 90 1550 ± 100 970 ± 150 620 ± 120

complex into products, and Γ is the surface concentration of hemin and is the number of electrons transferred (n = 2) [18,36]. The amount of water present in the solvent should have no influence on the hemin structure, but could significantly influence the rate of the reaction catalysed by hemin. To determine the optimal water content the catalytic activity was studied in ethanol solutions containing 1–10% v/v of added buffer (Fig. 4). No response was obtained in dry ethanol. The effect of added buffer on the response of the hemin sensor is shown in Table 1. The sensitivity of the sensor increased with the amount of added water, while the linear range also increased slightly with increasing buffer content. Imax and Kapp m values also increased with increasing amounts of added water in contrast to the trend obtained with HRP-Eastman electrodes, where the maximum values for Imax and Kapp m were obtained at ca. 4% added buffer [37,38]. As a result, a 95:5 v/v ratio of organic to aqueous content was selected for further experiments reflecting a compromise between a sufficiently high rate of catalysis and as low as possible water content. 3.4. Detection of hydrogen peroxide and kinetics of the catalytic reaction in nonaqueous solvents The response of the hemin sensor was examined in a range of solvents which contained 5% (v/v) or saturating amounts of aqueous buffer (TRIS, pH 7.0). Well-developed catalytic responses were obtained in primary alcohols (C1–C8), acetone, acetonitrile, ethyl acetate, tetrahydrofuran as well as in aqueous buffer. Some response was obtained in saturated toluene, however due to the negligible solubility of hydrogen peroxide in toluene, the catalytic activity of hemin could not be quantified. No catalytic response was obtained in DMF. The detection of hydrogen peroxide was carried out at an applied potential of 100 mV vs. Ag|AgCl. Normally, hydrogen peroxide is detected amperometrically at potentials above +0.6 V at unmodified platinum and carbon electrodes, respectively, potentials that are high enough for other compounds to be oxidised and interfere with the

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response. The catalytic detection of H2O2 (by e.g. hemin or peroxidases) is possible at low potentials is advantageous in diminishing the noise and the influence of electrochemically interfering substances [8,12]. In addition, the possibility of detecting hydrogen peroxide by heminmodified glassy carbon electrodes at potentials substantially more positive than the potential for Fe3+/Fe2+ conversion is indirect evidence, that hemin itself possesses some peroxidative activity and that similar intermediates to peroxidase compound I and compound II are probably involved in the catalytic cycle [10]. The response of hemin electrodes towards hydrogen peroxide is shown in Fig. 5. An initial linear part of the graph was in all cases followed by a region where the current levelled off. This may be a result of saturation by the substrate, hemin deactivation via oxidation by hydrogen peroxide [39], or most probably a combination of both factors. In the more polar solvents (methanol, ethanol, acetone and acetonitrile) the limiting currents were lower, with saturation and/or inactivation occurring at high peroxide concentrations. In less polar (more hydrophobic) solvents, the maximum catalytic current was attained at lower peroxide concentrations followed by a decrease in the current upon further addition of hydrogen peroxide (Fig. 5). For immobilised layers of enzymes on the electrode surface, four factors can influence the observed current [35]: (1) the rate of mass transport of the substrate to the electrode surface, (2) the rate of the reaction between the immobilised catalyst and the substrate, (3) substrate diffusion within a thick catalyst layer and (4) electron diffusion within the thick catalyst layer. As hemin forms a sub-monolayer film on the electrode surface, the rates of substrate and electron diffusion will not be rate determining. The catalytic current was independent of rotation rate between 500 and 2000 rpm, indicating that the kinetics of the overall reaction were not limited by mass transport of the substrate, but by the rate of the catalytic reaction. The rate of the catalytic reaction itself can be limited by the reaction of hemin with H2O2 or by the rate of electron transfer rate between hemin molecules and the electrode surface. As DET between hemin and the glassy carbon electrodes is fast, the reaction between hemin and H2O2 must be rate limiting. As mentioned previously, the Michaelis–Menten mechanism does not apply to peroxidases. This will also be the case for hemin, which will be inactivated by high peroxide concentrations prior to saturation. The inactivation of hemin would therefore be mainly responsible for the shape of the catalytic response (Fig. 5) where the current levels off much more rapidly than expected if it was just due to the saturation of the catalyst by the substrate. However, if the electrodes are not exposed to excessively high peroxide concentrations, they can be re-used with approximately 5% loss on re-use.

Fig. 5. Plot of response (current density per surface concentration of hemin) versus the concentration of hydrogen peroxide in nonaqueous solvents.

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Table 2 List of kinetic data for hemin-modified electrodes. Solvent

log P [46]

ε [87]

η (mN s m− 2) [44]

− Imax (A mol− 1)

Sensitivity (A mol− 1 mM− 1)

Kapp m (mM)

kapp cat (s− 1)

Imax / Kapp m (A mol− 1 mM− 1)

MeOH EtOH Propanol Butanol Pentanol Hexanol Octanol Aq. buffer Ethyl acetate ACN Acetone THF

− 0.77 − 0.31 0.25 0.90 1.40 2.03 3.00 − 1.38 0.73 − 0.34 − 0.24 0.46

33.62 25.00 22.20 17.80 16.90 13.30 11.30 78.30 6.11 37.50 20.70 11.60

0.676 1.209 2.522 2.948 4.650 6.203 10.64 0.891 0.473 0.397 0.391 0.550

7770 (440) 18950 (2320) 30930 (6940) 31280 (2770) 36650 (3200) 36770 (1850) 35710 (3630) 64960 (4890) 138300(14400) 19790 (1590) 6420 (580) 7200(150)

670 1220 2140 3020 3530 4360 11970 6340 20040 1130 440 1610

4.8 (3.0) 10.8 (6.5) 8.1 (4.8) 4.5 (1.0) 4.4 (0.4) 4.5 (0.7) 1.9 (0.5) 4.6 (0.7) 2.7 (1.2) 4.3 (1.7) 9.3 (2.6) 3.7 (1.4)

0.040 (0.002) 0.098 (0.012) 0.160 (0.036) 0.162 (0.014) 0.190 (0.017) 0.191 (0.010) 0.185 (0.019) 0.337 (0.025) 0.717 (0.074) 0.103 (0.008) 0.033 (0.003) 0.037 (0.001)

1620 1760 3820 6950 8330 8170 18800 14120 51220 4600 690 1950

The catalytic response was analysed in order to determine Kapp m and Imax. The values obtained from Lineweaver–Burk plots are sumapp marised in Table 2. The ratio of Imax / Kapp m (or kcat / Km ) provides a comparison of the catalytic efficiency of different systems [40]. An increase in Km can be indicative of diminished affinity of the catalyst for the substrate. The turnover number, kcat, can be obtained from: kcat = Imax = nFAC:

ð3Þ

The values of Imax obtained by Lineweaver–Burk plots are in all cases higher than the experimentally observed values of Imax (Table 2 and Fig. 5). Such differences may be a consequence of the inactivation of hemin at high concentrations of hydrogen peroxide, which would not be effectively reflected in the plots. The values of kcat are less than 1 s− 1 in all cases, and in most cases have values close to 0.1 s− 1 (Table 2). These values are much lower when compared to sensors constructed with whole enzymes as the biorecognition element, e.g.100 s− 1 reported for cytochrome c peroxidase [35] or 15 s− 1 for HRP [41]. Similarly the values of Kapp m are much higher for hemin-modified electrodes, ranging from 1–10 mM, compared to ca. 50 µM for immobilised cytochrome c peroxidase [35] and 0.2 mM for immobilised HRP [41], which may be indicative of lower affinity of hemin for hydrogen peroxide in comparison to the native enzymes. are compensated by the higher The lower values of kcat / Kapp m surface concentration of hemin compared to the lower coverages obtained with native enzymes [42], leading to higher catalytic currents and sensitivity for the hemin sensor. Hemin-modified electrodes displayed higher currents in aqueous buffer than in all nonaqueous solvents tested.

The catalytic efficiency of the hemin sensor (expressed as the ratio, Table 2) showed no dependence on the solvent Imax / Kapp m viscosity (1 / η) or the solvent dielectric constant (ε) (data not shown), similar trends were observed for HRP-Eastman AQ 55 sensor [43]. However, the catalytic efficiency was found to be linearly dependent on the hydrophobicity of the organic solvents used, (Fig. 6). Partitioning of the substrate between the solvent and the active site of the catalyst may be strongly affected by the presence of solvent molecules [40]. The rate of the catalytic reaction can also be affected by the substrate solubility in the solvent of interest, and by the activity of the catalyst in a given solvent. In the case of heminmodified electrodes the values of Kapp m were lower, and the catalytic efficiency higher with increasing hydrophobicity of the solvents (higher log P values) most likely due to preferable partitioning of H2O2 into the immobilised hemin layer [43]. This preferable partitioning was more pronounced in saturated solutions of 1-octanol (4% v/v) and ethyl acetate (3.5% v/v), where the polar H2O2 is forced to partition into the hemin layer due to very hydrophobic nature of these solvents, explaining the higher catalytic efficiencies observed in these solvents when compared to other 95% v/v solvent solutions. A comparison of the hemin sensor can be made to a mediated HRP system using ferrocenes as mediators [37,38], where the opposite trend was observed, i.e. the catalytic efficiency decreased with increasing solvent hydrophobicity. These findings were consistent with a previous report [45] on the oxidation of phenols by HRP in nonaqueous solvents. The decrease in catalytic efficiency could be explained by a destabilisation of the transition-state complex formed between the mediator and the enzyme in hydrophobic solvents. In all solvents tested the catalytic efficiency decreased with increasing solvent hydrophobicity due to diminished partitioning of hydrophobic substrates (phenols) to the HRP active site. Larger concentrations of the substrates were therefore necessary to saturate the enzyme, leading to increased values of apparent Km, which was also observed for the HRP-Eastman-based sensor [38]. The hemin sensor described here utilises a polar substrate. Partitioning of peroxide into the hemin layer will become more favourable as the solvent becomes more hydrophobic, accounting for the trend of increasing catalytic efficiency with increasing solvent hydrophobicity. 4. Conclusions

Fig. 6. Plot of catalytic efficiency of hemin electrodes in the catalytic reduction of H2O2 versus solvent hydrophobicity (log P).

The catalytic activity of hemin adsorbed on graphite for the detection of hydrogen peroxide in aqueous and organic solvents has been described. Immobilised hemin displayed reasonable stability in aqueous buffer, organic solvents and both aqueous and organic solutions of hydrogen peroxide. In all solvents tested the response of the sensor was independent of the rotation rate suggesting that the kinetics of reduction of hydrogen peroxide were limited by the rate of the catalytic reaction between hemin and hydrogen peroxide rather than by the mass transport of hydrogen peroxide to the electrode surface. The kinetic

Z. Brusova, E. Magner / Bioelectrochemistry 76 (2009) 63–69 app characteristics (Imax, sensitivity, Kapp m , kcat, Imax /Km ) were evaluated in eleven organic solvents containing 5% v/v or saturating amounts of aqueous buffer. A linear correlation (R2 = 0.91) was observed between the catalytic efficiency (Imax /Kapp m ) and the solvent hydrophobicity (log P) in all solvents. Preferable partitioning of hydrogen peroxide to the polar hemin layer in more hydrophobic solvents can account for the trend observed.

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