Fenton-like Inactivation of Tobacco Peroxidase Electrocatalysis at

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Fenton-Like Inactivation of Tobacco Peroxidase Electrocatalysis at Negative Potentials Jose Luis Olloqui-Sariego, Galina S. Zakharova, Andrey A. Poloznikov, Juan Jose Calvente, Dmitry M. Hushpulian, Lo Gorton, and Rafael Andreu ACS Catal., Just Accepted Manuscript • DOI: 10.1021/acscatal.6b01839 • Publication Date (Web): 28 Sep 2016 Downloaded from http://pubs.acs.org on October 3, 2016

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Fenton-Like Inactivation of Tobacco Peroxidase Electrocatalysis at Negative Potentials José Luis Olloqui-Sariego,a Galina S. Zakharova,b Andrey A. Poloznikov,b Juan José Calvente,a Dmitry M. Hushpulian,b Lo Gorton,c Rafael Andreu a,* a

Department of Physical Chemistry. University of Sevilla. Profesor García González 1. 41012

Sevilla. Spain. b

D. Rogachev center of Pediatric Hematology, Oncology and Immunology, 1 Samory Mashela

str., Moscow 117997, Russia. c

Department of Biochemistry and Structural Biology. Lund University, P. O. Box 124, 221 00,

Lund, Sweden. Corresponding Author * [email protected] Tlf: (0034)-955421002

ABSTRACT: The effect of the operational potential on the stability of electrochemical biosensors is particularly relevant in the case of peroxidase biosensors, since these enzymes can catalyze the reduction of hydrogen peroxide via either a high-potential redox cycle (involving Compound I, Compound II and Fe(III)), or a low-potential redox cycle (involving Fe(III) and Fe(II)). Herein, it is shown that recombinant Tobacco Peroxidase immobilized on a graphite

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surface displays two well separated electrocatalytic waves, associated with each of these two catalytic cycles. While continuous scanning in the high-potential region does not alter significantly the electrocatalytic current, it is shown that just modest incursions into the lowpotential region cause an irreversible loss of the electrocatalytic response. A quantitative analysis of the inactivation extent as a function of time, potential and hydrogen peroxide concentration is shown to be consistent with a fast inactivation brought about by hydroxyl radicals generated by a Fenton-like mechanism. Accordingly, the inactivation process is shown to slow down by addition of radical scavengers to the solution. Preliminary results indicate that the same inactivation process may also be present in horseradish peroxidase modified electrodes.

TOC GRAPHICS

KEYWORDS Tobacco Peroxidase; Bioelectrocatalysis; Direct electron transfer; Hydroxyl Radical; Fenton Reaction.


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Introduction Direct electron transfer between redox enzymes and electrodes has extensively been studied to gather fundamental information on enzymatic processes,1 and to develop new catalytic structures for their application in biofuel cells2 and biosensors.3 It is generally accepted that to obtain adequate electrocatalytic currents, immobilized enzymes should preserve their native structure and be adequately oriented to facilitate a fast electronic exchange with the electrode.4 However, much less attention has been paid to the effect of the operational potential on the stability of these enzymes. This effect is expected to be particularly relevant in the case of peroxidase-based electrochemical biosensors, because peroxidases can catalyze the reduction of hydrogen peroxide via either the Compound I/Compound II/Fe(III) high-potential redox cycle, or the Fe(III)/Fe(II) low-potential redox cycle.5-8 The former operates at potentials close to the formal potentials of the two redox couples involved (i. e. ∼+0.9 V vs. NHE at pH 7),9 and it requires a precise location of some histidine and arginine residues around the heme group to stabilize the Compound I and Compound II oxyferryl states and to facilitate the associated proton exchange.10 The low-potential redox cycle operates at more negative potentials (typically, ≤ 0 V vs. NHE at pH 7),9 and it does not require a specific amino acid disposition around the heme group, since both partially denatured cytochrome c11 and bare hemin12,13 have been shown to act as electrocatalysts of H2O2 reduction under anaerobic conditions. While there is a broad consensus on the mechanistic details of the high-potential cycle,14 two alternative mechanisms have been proposed for the low-potential cycle.11 The first assumes that H2O2 and Fe(II) react initially to produce Fe(IV)-oxyferryl (i.e. Compound II in the high-potential cycle), which accepts then two electrons from the electrode, generating thus the electrocatalytic current and restoring the initial Fe(II) state.15,16 The second mechanism corresponds to the so called electro-


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Fenton reaction,17,18 where Fe(II) reduces H2O2 to produce a hydroxyl radical and Fe(III), which is then reduced back by the electrode to Fe(II) to restart the cycle. It may be worth stressing that this last mechanism implies the continuous formation of hydroxyl radicals, whose high and unselective reactivity19 is likely to reduce drastically the operating life of the biosensor. In fact, herein we show how a quantitative analysis of the catalytic current inhibition as a function of the applied potential, time and hydrogen peroxide concentration is consistent with a fast enzymatic inactivation at negative potentials due to hydroxyl radicals generated at the electrode surface.

Materials and Methods

Enzyme production Wild-type recombinant TOP (r-TOP, hydrogen peroxide oxidoreductase, EC 1.11.1.7) was expressed in the form of inclusion bodies in E.coli BL21(DE3) CodonPlus cells as was described earlier.20 The construction of the expression vector and the procedure of r-TOP expression, refolding and purification are covered in detail elsewhere.20 Briefly, biomass from 600 mL of culture medium was disrupted and the precipitate of inclusion bodies was washed, solubilized in 6 M urea and added drop by drop to the refolding medium, containing 1.8 M urea, 0.1 mM DTT, 0.5 mM oxidized glutathione, 3 mM СаСl2, 5 µM hemin, 5% glycerol in 50 mM Tris-HCl pH 9.5. After in vitro reactivation r-TOP was concentrated and purified on a Toyopearl HW-55 column. The concentration of peroxidase was determined from the absorbance at 403 nm, using the experimentally determined extinction coefficient value of 108.0 ± 0.5 mM-1cm-1 for the protein monomer.21 The resulting preparation was homogeneous as judged by SDS-PAGE. The activity of purified r-TOP towards ABTS was about 4100 U/mg and the RZ value was 3.0. Reagents and Chemicals HRP isoenzyme C (EC 1.11.1.7, activity 1067 U mg-1) from Fluka was used as received. 1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC), N-hydroxysulfosuccinimide sodium salt (NHS), hydrogen peroxide, 2,2’-azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS)


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from Sigma Aldrich and glucose from Panreac were high purity reagents and were used as received. Sodium phosphate buffer solutions (SPB) were prepared from dihydrogen sodium phosphate (from Fluka) and water purified with a Millipore Milli-Q system (resistivity 18 MΩ cm). Enzyme immobilization Pyrolytic graphite electrodes were constructed by fitting a rod of highly oriented pyrolytic graphite from Mineral Technologies into a PEEK casing, so that it exposed the edge of the graphite planes with a circular geometric area of 0.07 cm2. Prior to enzyme immobilization, graphite electrodes were polished with abrasive P2400 sandpaper, then they were rinsed with Millipore water and dried with an argon stream. Covalent coupling was carried out by exposing the electrode surface for 20 h (at 4ºC) to a mixture of three droplets: a 4.5 µL drop of 20 mM NHS, a 5.5 µL drop of 40 mM EDC and a 7 µL drop of a 0.26 mg mL-1 r-TOP solution, these solutions were also 0.01 M SPB at pH 6. Then, the electrodes were thoroughly rinsed with water and washed with the working buffer solution. Electrochemical measurements Electrochemical measurements were performed with an AUTOLAB PGSTAT 30 potentiostat, from Eco Chemie B.V, in a conventional three-electrode undivided glass cell, equipped with a gas inlet and thermostated with a water jacket. The counter and reference electrodes were a Pt bar and an Ag|AgCl|NaCl saturated electrode, respectively. The reference electrode was connected to the cell solution via a salt bridge, and kept at room temperature (23 ± 2ºC) in a nonisothermal configuration. Reported potential values have been corrected to the SHE potential scale by adding +192 mV to the experimental potential values. The rotating disk electrode was operated at 1000 r.p.m. with an AMSFRX analytical rotator from Pine. All measurements were carried out under an argon atmosphere. To minimize any enzyme inactivation by H2O2 (see Figure S4 in ref. 22), all measurements were carried out at 0ºC. Working solutions contained 20 mM sodium phosphate buffer at pH 7.0.


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Results and discussion General electrocatalytic behavior Though a variety of peroxidase-based biosensors have been described before,3 only the electrocatalytic response associated with either the high-potential23,24 or the low-potential25,26 cycle was reported in each case, thereby precluding a direct assessment of their distinct effect on the stability of a given biosensor. However, we have shown recently22 that the simultaneous incubation of recombinant Tobacco Peroxidase (r-TOP) with amide cross-coupling reagents at a pyrolytic graphite electrode leads to the covalent immobilization of a mixture of denatured and native enzymes electronically wired to the electrode surface. Each group of these electroactive enzymes is expected to adopt a different route to catalyze the electroreduction of H2O2. In agreement with this hypothesis, the voltammetric response of a graphite electrode modified according to the former incubation protocol shows two electrocatalytic waves, as illustrated in Figure 1. In the absence of H2O2, a single and reversible surface wave with an average peak potential of -0.100 V vs NHE is observed, and it is associated with the Fe(III)/Fe(II) redox conversion of any r-TOP enzyme that exchanges electrons with the electrode (Figure 1a). The non-turnover signal associated with the more anodic electrocatalytic wave cannot be observed in the absence of H2O2, since the formation of Compound I requires the reaction of the Fe(III) enzyme with H2O2. Upon addition of H2O2 to the solution, two well separated waves are obtained at ca. 0 V and +0.7 V (denoted as LP and HP in Figure 1b, respectively). Their proximity to the Fe(III)/Fe(II) and to the Compound I/Compound II/Fe(III) redox potentials9 indicates that these two waves are generated by (denatured) enzymes operating through the lowpotential cycle and by (intact) enzymes operating through the high-potential cycle, respectively.


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Figure 1. Voltammetric response of r-TOP immobilized on a graphite electrode. (a) In the absence of H2O2 recorded at 2 V s-1. (b) In the presence of 160 µM H2O2 recorded at 0.02 V s-1. (c) In the presence of 120 µM H2O2, before (red curve) and after three successive 5 min immersions (green, pink and blue curves, respectively) in a 9 M urea aqueous solution. (d) Influence of the cathodic vertex potential on the voltammetric response after recording ten consecutive cyclic voltammograms at 0.02 V s-1 in the presence of 240 µM H2O2. First scan is depicted in red and tenth scan in green. Voltammograms at the more positive potentials have been shifted vertically for clarity. Dashed grey voltammograms in (c) and (d) were recorded in the absence of H2O2. Other experimental conditions: 0.02 M SPB, pH 7 and 0ºC.

This assignment is further confirmed by observing that the HP and LP signals decrease and increase, respectively, when the modified electrode is put briefly into contact with a concentrated urea solution (see Figure 1c). This urea solution is expected to induce the conversion of native enzymes into their denatured form, so that the behavior observed in Figure 1c is consistent with


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our previous assumptions on the origin of the two catalytic signals. A more detailed analysis shows that the area loss of the HP signal in the presence of urea coincides with the area increase of the LP signal, and it amounts to one fifth of the total LP area, thereby suggesting that ∼ 20% of the heme groups that are wired to the electrode retain their native structure and contribute to the HP catalysis. Since the two HP and LP electrocatalytic currents increase with H2O2 concentration (see Figure S2), both could in principle be used to quantify the H2O2 content in solution. However, their effects on the biosensor stability are markedly different. Figure 1d illustrates how it suffices to perform ten consecutive scans down to ca. -0.3 V to suppress irreversibly all the electrocatalytic activity of the electrode, while continuous scanning at more positive potentials does not modify the electrocatalytic response (see also Figure S3). Therefore, the loss of enzymatic activity can be traced back to the simultaneous presence of both Fe(II) and H2O2 at the electrode surface. This requirement should not be too surprising, since these two species are known to react in solution yielding a strong oxidizing product, whose chemical identity has been debated for a long time.27,28 Fenton-like inactivation mechanism and kinetics The two main Fe(II) oxidation reactions that have been proposed are: 11,29 ox ,1 a Enz ( Fe(II) ) + H 2O2  → Enz ( Fe(IV)=O ) + H 2O

k

ox ,1b Enz ( Fe(II) ) + H 2O2  → Enz ( Fe(III) ) + OH − + • OH

k

(1a)

(1b)


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where Enz ( Fe(II) ) , Enz ( Fe(III) ) and Enz ( Fe(IV)=O ) stand for the ferrous, ferric and Compound II forms of r-TOP enzymes, respectively, and kox,1a and kox,1b are the rate constants for the bi- and mono-electronic oxidations of Fe(II) according to reactions 1a and 1b, respectively. These two reactions lead to equivalent kinetic expressions, but they do not predict the same inactivation extent for native (Enznat) and denatured (Enzdenat) enzymes. In fact, native enzymes that are oxidized according to eq. 1a are expected to remain intact, since they have been shown to produce Compound II at more positive potentials without any loss of catalytic activity. Therefore, to achieve a complete inactivation of both native and denatured enzymes, as observed in Figure 1b (see also Figure S1), at least some denatured or native enzymes should be oxidized according to eq. 1b, so that the mobile • OH radicals would be able to oxidize any nearby TOP molecules (Enz) according to: kinac Enz + • OH  → Inactive products

(2)

where the inactivation rate constant kinac is expected to be much larger than either kox,1a or kox,1b.19 Since denatured r-TOP enzymes do not meet the structural requirements to generate HP electrocatalytic currents and, therefore, are not likely to produce Compound II according to eq. 1a, we will assume that they are oxidized according to eq. 1b. Since the inactivation process takes place at rather negative potentials, where Compound I and Compound II are quickly reduced to either Enz(Fe(III)) or Enz(Fe(II)), their steady-state concentrations can be neglected, and the total surface concentration of electroactive enzymes can be approximated by the sum of the surface concentrations of their ferric and ferrous forms: T T T ΓEnz = ΓEnz + ΓEnz ( Fe( III ) ) ( Fe( II ) )

(3)


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where the surface concentration of each redox state: T nat denat ΓEnz = ΓEnz + ΓEnz , i=Fe ( II ) ,Fe ( III ) (i ) (i ) (i )

(4)

nat denat is the sum of the contributions from the native ( ΓEnz ) and denatured ( ΓEnz ) populations of (i ) (i )

electroactive enzymes The rate of enzyme inactivation νinac is assumed to be the same for native and denatured enzymes, and can be expressed as:

νinac =

T dΓEnz T = − kinac ΓEnz c•OH ( 0,t ) dt

(5)

where c•OH ( 0,t ) is the local concentration of hydroxyl radicals at the electrode surface. dc •OH ( 0 ,t ) dt

denat T = koxdenat ΓEnz ( Fe( II ) ) cH 2O2 ( 0 ,t ) − kinac ΓEnz c • OH ( 0 ,t ) ≈ 0

(6)

where cH 2O2 ( 0,t ) is the local concentration of hydrogen peroxide at the electrode surface, and denat k oxdenat is the rate constant for radical formation according to eq. 1b (i.e. kox = kox ,1b ).

Since hydroxyl radicals are very reactive species (typical k inac values fall within the 108 M-1s-1 < kinac < 1010 M-1s-1 range19), it seems reasonable to assume that their concentration obeys the steady-state hypothesis, dc OH / dt ≈ 0 , so that the following expression is obtained from eq. 6: denat koxdenat ΓEnz ( Fe( II ) ) cH 2O2 ( 0 ,t ) c •OH ( 0,t ) = T kinac ΓEnz

(7)


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At a given potential Einac where inactivation takes place, the surface concentrations of the ferric and ferrous forms of the denatured enzyme are related through Nernst equation:

Einac = E

0 denat

denat RT ΓEnz ( Fe( III )) + ln denat nF ΓEnz( Fe( II ))

(8)

0 where Edenat is the formal potential of the Enzdenat(Fe(III))/Enzdenat(Fe(II)) redox couple, which is

assumed to take the same value for the native and denatured electroactive enzymes, in agreement with the simple voltammetric behavior observed in the absence of H2O2 (see Figure 1a). Then, denat denat denat = ΓEnz + ΓEnz by combining eq. 8 and ΓEnz , the surface concentration of the ferrous form ( Fe( II ) ) ( Fe( III ))

can be expressed in terms of the total surface concentration of denatured electroactive enzymes denat ΓEnz and the applied potential Einac as:

denat ΓEnz ( Fe( II ) ) =

denat ΓEnz 1 + ξinac

(9)

0 where ξinac = exp ( Einac − Edenat ) F / RT  . Then, eqs. 7 and 9 can be combined to obtain:

c •OH ( 0 ,t ) =

koxdenat χ denat cH 2O2 ( 0 ,t ) kinac (1 + ξinac )

(10)

denat T where χ denat = ΓEnz / ΓEnz is the fraction of electroactive enzymes that are denatured. By inserting

this expression for c•OH ( 0,t ) into eq. 5, we obtain the rate of inactivation as a function of the inactivation potential and the concentrations of wired enzymes and hydrogen peroxide:


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νinac

T T koxdenat χ denat cH2O2 ( 0,t ) ΓEnz dΓEnz = =− dt 1 + ξinac

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(11)

Under the steady-state conditions prevailing in rotating disk experiments, the concentration profile of hydrogen peroxide becomes time-independent, and its concentration at the electrode surface can be expressed as:

 i cH2O2 ( 0 ) = cH∗ 2O2 1 −   iL 

(12)

where cH∗ O is the hydrogen peroxide bulk concentration, i is the steady state current and iL is the 2 2

diffusion-convection limiting current given by the Levich equation: − iL = 0.62ν1/ 6 DH2 /23O2 ω1/ 2 c*H 2O2 2 FA

(13)

where DH2O2 , ν, and ω are the hydrogen peroxide diffusion coefficient, the kinematic viscosity of the solution, and the electrode rotation rate, respectively. Under our experimental conditions * * i / iL ≤ 0.01 , so that cH2O2 ( 0) ≈ cH2O2 in eq. 12, and cH2O2 ( 0,t ) can be replaced by cH2O2 in eq. 11 to

a good approximation. Then, eq. 11 can be integrated between inactivation times 0 and tinac , T T ,0 where the surface concentrations of active enzymes are ΓEnz and ΓEnz , respectively, to obtain at a

given inactivation potential: T koxdenat χ denat c*H 2O2 ΓEnz ln T ,0 = − tinac ΓEnz 1 + ξ inac

(14)


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Figure 2. (a) Scheme of the potentiostatic pulse experiments performed at the rotating disk electrode. First, the electrocatalytic current (i)tinac=0 of a freshly modified electrode is measured at Emeas = 0.30 V, where TOP inactivation does not occur. Then, the potential is stepped to a potential Einac within the 0.14 V ≥ Einac ≥ -0.31 V range, where inactivation takes place for a time tinac. Finally, the electrocatalytic current i is measured again at 0.30 V. (b) Influence of the inactivation potential on i. (c) Influence of hydrogen peroxide concentration and inactivation time on i. Symbols in (b) and (c) are experimental 0 values and solid lines have been computed from eq 4 with E denat = -0.1 V and koxdenat ,ap = 150 M-1s-1. Other

experimental conditions: 0.02 M PBS, pH 7, 0 ºC. Error bars indicate the standard deviation of at least three replicated measurements for each data point. Average standard errors in Figures 2b and 2c are ±0.03.

To assess the kinetics of the inactivation process, the time evolution of the population of native enzymes was sampled in an experiment involving two steps. First, inactivation of a freshly modified electrode was allowed to proceed at a potential Einac for a time tinac , so that the number T T ,0 of electroactive enzymes was reduced from ΓEnz to ΓEnz . Then, the potential was stepped to a

potential Emeas = 0.3 V where no further inactivation takes place, and where the recorded


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electrocatalytic current (i) is proportional to the remaining surface concentration of native enzymes, i.e. i / ( i )t

inac = 0

nat nat ,0 (the validity of eq 14 was checked independently by = ΓEnz / ΓEnz

determining spectrophotometrically the catalytic activity of a TOP modified electrode, after it was subjected to an inactivating pulse potential, see Supporting Information). In agreement with our previous assumption that inactivation of native and denatured enzymes takes place at the nat nat ,0 denat denat ,0 T T ,0 same rate, i.e. that ΓEnz , eq. 14 can then be rearranged as: / ΓEnz = ΓEnz / ΓEnz = ΓEnz / ΓEnz

ln

i

(i )

tinac = 0

=−

koxdenat χ denat c*H 2O2 1 + ξinac

tinac = −

koxdenat ,ap c*H 2O2 1 + ξinac

tinac

(15)

where koxdenat ,ap = koxdenat χ denat . This expression is easily amenable to experimental verification, as illustrated in Figure S4 (Supporting Information). As it may be observed in Figure 2, eq. 15 describes satisfactorily the variation of the inactivation ratio i / ( i )t

inac = 0

with time, potential and hydrogen peroxide concentration with only

one adjustable parameter value koxdenat ,ap = 150±10 M-1s-1. This value represents a lower limit for the true oxidation rate constant koxdenat , since χ denat ≤ 1. If we adopt χ denat = 0.8, as derived before from the ratio of the HP and LP voltammetric areas, then koxdenat = 190 M-1s-1. We are not aware of analogous rate constants reported for other peroxidases, but this koxdenat value is significantly larger than 47 M-1 s-1 determined for cytochrome C-Fe(II) oxidation by H2O2 in homogeneous solution at 25ºC.30 It seems likely that the faster Fenton kinetics associated with our modified electrode reflect an easier access of H2O2 to the porphyrin ring in denatured r-TOP molecules, and possibly also some favorable changes in the coordination and local environment of the iron center.31,32 Besides its overall success, our kinetic model tends to overestimate the inactivation


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extent at the highest cH∗ O and longest tinac values (see Figure 2 c), this trend is likely to reflect 2 2

the reduced number of hydroxyl radicals that are being produced under these circumstances and their difficulty to reach and inactivate the remaining native enzymes.

Figure 3. Catalytic cyclic voltammograms of r-TOP immobilized on graphite. (left) In the absence of glucose, (red curve) first scan after adding 240 µM H2O2, (green curve) tenth scan after H2O2 addition. (right) In the presence of 2.5% (m/v) glucose in solution, (red curve) first scan after adding 240 µM H2O2, (green curve) tenth scan after H2O2 addition. Inset Plot: Dependence of the inactivation ratio at 0.3 V on the glucose concentration in solution. Grey dashed voltammograms were recorded before adding H2O2 to the solution, other experimental conditions: 0.02 M PBS, pH 7, 0.02 Vs-1and 0 C.

In order to corroborate the involvement of free radicals in the inactivation of r-TOP, we explored the effect of well-known hydroxyl radical scavengers,33,34 such as glucose, polyethylene glycol and bovine serum albumin, on the stability of the electrocatalytic response. Figure 3 illustrates how the presence of glucose in solution helps to preserve the electrocatalytic current


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(we have recorded similar, though somewhat less marked, protective effects in the presence of polyethylene glycol and bovine serum albumin in solution). It may also be observed how the inactivation ratio increases with glucose concentration until it reaches a plateau, as it is to be expected when adsorbed glucose molecules are responsible of the radical scavenging effect. However, it should be pointed out that the presence of radical scavengers slows down, but it does not stop the inactivation process.

Figure 4. Catalytic cyclic voltammograms of wild HRP-C immobilized on graphite. Influence of the cathodic potential vertex on the voltammetric response after recording twenty consecutive cyclic voltammograms in the presence of 240µM H2O2. Voltammograms recorded at more positive potentials have been shifted vertically for clarity. Other experimental conditions as in Figure 3.

In summary, we have shown in this work how surface immobilized r-TOP molecules become inactivated at potentials where their iron centers are reduced to Fe(II) in the presence of H2O2. It is then fair to wonder whether the same inactivation process applies to wild HRP, which


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constitutes the basic building block of most peroxidase based biosensors. Figure 4 shows some preliminary results obtained with HRP isoenzyme C, which illustrate the influence of the scanning potential region on the stability of the electrocatalytic response. They were recorded under the same conditions as those in Figure 1d corresponding to r-TOP, allowing thus for a direct comparison between the two sets of results. This comparison shows clearly that the same inactivation process is also present for the HRP modified electrode, though it appears to evolve at a slower pace. ASSOCIATED CONTENT Supporting Information. Non-turnover voltammograms recorded before and after enzyme inactivation. Influence of the cathodic vertex potential on the calibration curves of freshly prepared r-TOP modified graphite electrodes. Effect of the cathodic vertex potential on the electrocatalytic inactivation of immobilized r-TOP. Experimental verification of linearized eq. 15. Spectrophotometric assessment of the potential-induced inactivation process. This material is available free of charge via the Internet at http://pubs.acs.org. Notes The authors declare no competing financial interests.

ACKNOWLEDGMENT J. L. O., J. J. C. and R. A. acknowledge financial support from the Spanish Ministry of Economy and Competitiveness and the European Union FEDER (grants CTQ2014-52641-P and CTQ2015-71955-REDT (ELECTROBIONET) ) and L. G. from the Swedish Research Council (grant 2014-5908).


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TOC 271x262mm (88 x 88 DPI)


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Figure 1 103x105mm (300 x 300 DPI)


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Figure 2 107x66mm (300 x 300 DPI)


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Figure 3 91x92mm (300 x 300 DPI)


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Figure 4 88x92mm (300 x 300 DPI)


Fenton-like Inactivation of Tobacco Peroxidase Electrocatalysis at

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