Electrochemical approach for monitoring the catalytic action of immobilized catalase

Angel Peshkov; Merin Shukri; Mariya Pimpilova; Ilia Iliev; Nina Dimcheva

doi:10.1515/chem-2025-0143

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Electrochemical approach for monitoring the catalytic action of immobilized catalase

Angel Peshkov , Merin Shukri , Mariya Pimpilova , Ilia Iliev und Nina Dimcheva

Veröffentlicht/Copyright: 29. März 2025

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Abstract

Heterogeneous enzymatic activity of fungal catalase from Penicillium chrysogenum in the hydrogen peroxide (H₂O₂) disproportionation reaction in a neutral aqueous buffer solution has been examined by means of constant-potential chronoamperometry. A recently developed catalytic peroxide electrode has been used as both an electrochemical detector and a high-speed low-noise stirrer. The determination of immobilized enzyme activity has been performed in an electrochemical system containing a sample of immobilized fungal catalase (from P. chrysogenum) and a rotating catalytic electrode sensitive to H₂O₂ concentration placed at a close distance to the sample. Heterogeneous catalytic activities of fungal catalase for H₂O₂ decomposition have also been assayed in the presence of 0.5 and 1.0% ethanol and methanol. Alterations of catalase catalytic action have been found in the presence of alcohols: while in the absence of alcohols its kinetics obeys the Michaelis–Menten mechanism, in the presence of alcohols it is described by Hill’s model. The apparent kinetic constants of the enzyme-catalyzed process have been proven to differ depending on the type of alcohol and its amount in the medium. The obtained results imply that the use of the electrocatalytic approach can be considered a promising alternative for kinetic characterization of immobilized enzymes, provided that the enzyme substrate is electrochemically active.

Keywords: immobilized catalase; enzyme kinetics; alcohols; peroxidase-like activity; electrochemical method

1 Introduction

Catalase is an enzyme found in all aerobic living organisms, including microorganisms, plants, and animals. It usually exists as a homotetramer composed of four identical polypeptide chains (subunits), each containing hundreds of amino acid residues.

There are differences in catalase structure, which reflect the diverse roles and the adaptation models of enzyme in different organisms: catalases isolated from ascomycetes often have larger subunits compared to catalases of animal origin. For example, fungal catalases can have subunit sizes ranging from 75 to 84 kDa, while mammalian ones typically have smaller subunits [1]. Catalases are metalloproteins containing iron-heme groups, enabling the enzyme to catalytically decompose H₂O₂, and the types of heme groups may differ from species to species – while bacterial and fungal catalases often have heme d as a prosthetic group, the enzymes from animal origin usually comprise heme b [1]. Moreover, mammalian catalases contain NADPH as an enzyme cofactor [2] that protects it from inactivation and helps the regeneration of the enzyme in its active form, unlike catalases from plant or fungal origin.

Catalase is the first discovered enzyme from the antioxidant system of living cells, the physiological function of which is to decompose hydrogen peroxide (H₂O₂) generated during metabolic processes to water and oxygen. Changes in its activity in plants are often associated with plant stress responses [3] arising from the changes in environmental conditions or biotic challenges [4,5]. In humans, catalase functions as a detoxifying enzyme, which oxidizes nitrites and alcohols [1] coming from cellular respiration, and its activity in blood and tissues might be taken as a reliable marker of metabolic disorders [6] since the enzyme is a part of the alcohol-metabolizing microsomal system [7].

Catalase is an ultrafast catalyst that possesses one of the highest turnover numbers [8], which predetermines its wide spectrum of applications. In medical diagnostics, lowered catalase activity is a biomarker for diabetes, liver diseases [7], some types of cancer [1], or age-associated degenerative diseases [9]. Catalase tests are widely employed in microbiology as an instrument for the discrimination between aerobic and anaerobic microorganisms [10]. The enzyme activity assays are used to monitor the safety of food products, as they facilitate spotting of microbial contamination, and hence ensure proper functioning of food preservation processes. These assays find prospective applications in environmental sciences as well, as they are used to assess the oxidative stress in plants and other organisms exposed to pollutants [3,4,5], this way helping in understanding the impact of environmental stressors on living organisms.

Catalase finds applications in various industrial biotechnological processes in food industries, such as the removal of residual H₂O₂ from raw milk, which undergoes cold pasteurization prior to cheese production [11,12].

Catalase activity assays are used in basic and applied research to study oxidative stress, aging, and the role of antioxidants in protecting cells from damage [9,13]. In addition to its function as H₂O₂₋ and nitrite-detoxifying enzymes, catalase possesses a complex role in alcohol metabolism [14] during which reactive oxygen species (ROS) can be produced, thus affecting both cellular functions and enzymatic activity. Together with alcohol dehydrogenase and cytochrome P450, the enzyme oxidizes ethanol to acetaldehyde, however, using H₂O₂ as an oxidizer, with a K_M value of 12 mM [15].

Some fungal catalases may also have the ability to oxidize hydrogen donors [16] taken at very low concentrations using H₂O₂ as an oxidizing agent, and this peroxidase-like activity of immobilized catalase in the oxidation of phenolic compounds has been explored earlier [17].

In addition to the standard method for the catalase activity assay, where H₂O₂ concentration decay is monitored spectrophotometrically at 240 nm, several different variations are reported, employing spectrophotometric measurements of the absorbance of colored complexes formed in the presence of H₂O₂ [18,19,20]. Another approach to quantify the activity of the enzyme is to determine the concentration of oxygen generated upon H₂O₂ decomposition by means of an oxygen-sensitive electrode (oximetry) [21,22].

The aim of the present work is to demonstrate the applicability of electrochemical methods in the determination of the activity of immobilized catalase. Having in mind that in living cells catalase is confined in cell organelles such as peroxisomes or mitochondria [23], enzyme immobilization on a solid support mimics to some extent enzyme’s natural state in living tissues. Moreover, catalase immobilization permits its easy separation from the reaction medium. To monitor its catalytic function, a bioreactor consisting of a conventional electrochemical cell with a peroxide-sensitive indicator electrode and immobilized fungal catalase on a glass support placed opposite to it at a close distance was proposed. This bioreactor arrangement allowed the evaluation of the immobilized enzyme catalytic activity under hydrodynamic conditions both in the absence and presence of low concentrations of methanol and ethanol.

Experimental results showed that the chosen approach is viable not only for the determination of enzymatic activity but also for monitoring alteration of the enzyme kinetics from Michaelis-type to Hills’ type observable in the presence of alcohols.

2 Experimental

2.1 Materials

Catalase (EC 1.11.1.6) from the strain Penicillium chrysogenum 245 (registration number 677 in NBIMCC, Bulgaria) with specific activity ≥ 250 kU/g, isolated and purified following the biotechnology process proposed by Prof. Ilia Iliev in the form of a deep green powder, has been used as the biological material in this study. Catalase purification was performed, as reported by Sutay Kocabas et al. [24].

In electrochemical studies, the working electrodes were discs from glassy carbon with the diameter of the working surface d = 2 mm (Metrohm, Utrecht, The Netherlands); H₂O₂ (Merck), tert-butyl hydroperoxide (Acros Organics), Nafion™ 117, 5% suspension in water–alcohol mixture (Sigma-Aldrich), K₂HPO₄, and KH₂PO₄ (Acros) were of analytical grade and used without further purification. Anhydrous solvents – methanol and ethanol – were of HPLC grade (Fisher) and used as received. All other chemicals were of reagent grade and used as received.

Buffer solutions with pH = 7.00 ± 0.05, typically 0.1 M concentration, were prepared with monobasic and dibasic potassium phosphate dissolved in ultrapure water (0.055 µS cm⁻¹, total carbon ≤ 2 ppb, B30 Adrona Bio, Vilnius, Lithuania). The buffer solutions were adjusted to the desired pH with a pH meter FiveEasy (Mettler-Toledo, OH, USA).

2.2 Apparatus and measurements

All electrochemical measurements were performed in a conventional three-electrode cell (working volume: 20–100 ml, Metrohm, Utrecht, The Netherlands), with catalyst-modified discs of glassy carbon as the working electrode, Ag|AgCl (sat. KCl) as the reference electrode, and platinum foil as the auxiliary electrode. An electrochemical workstation Autolab 302 N (Metrohm, Utrecht, The Netherlands) with Nova 2.1.6 software was used in all experiments.

Chronoamperometric measurements were performed under constant stirring, using the rotating disk electrode (RDE) module at a rotating speed between 500 and 3,000 rpm.

When necessary, the working medium was deaerated with argon gas (99.99% purity).

Electron microscopy was performed with a JSM 6390 (JEOL, Japan) scanning electron microscope equipped with an energy-dispersive X-ray (EDX) spectrometer.

2.3 Preparation of the working electrode

The peroxide-sensitive electrode, used as an electrochemical detector, was prepared as described earlier [25]. In brief, the glassy carbon electrode (GCE) surface was cleaned by polishing with 0.05 µm suspension of γ-Al₂O₃ and rinsed with ultrapure water. The polishing step was followed by ultrasonication in distilled water for 3 min and then dried at room temperature. Next, a 5 µl drop of the catalyst containing suspension was cast onto the GCE surface and dried at room temperature for 16 h. The modifying phase was prepared through dispersion of 2 mg of catalyst (Co–g-C₃N₄) in 1 mL of the polymer suspension of Nafion assisted by ultrasonication for at least 45 min.

2.4 Catalase immobilization

For the examination of immobilized catalase activity, clean glass substrates for optical microscopy with dimensions approx. 1 cm × 2 cm were used. Before use, the glass substrates were sonicated in 95% ethanol for 3 min and then rinsed and sonicated in ultrapure water twice for 2 min each time. On thoroughly cleaned glass plates, the enzyme was immobilized using two alternative binding strategies:

about 50 μL of catalase solution with a concentration of 5 mg/mL was drop-cast on the glass, 25 μL of 0.5 mM glutaric aldehyde aqueous solution was added, mixed with the pipette tip, and allowed to crosslink for 1 h in the refrigerator;
about 50 μL of catalase solution with a concentration of 5 mg/mL was drop-cast on the glass, 25 μL of 0.2% aqueous suspension of the ionomer Nafion was added, mixed with the enzyme solution, and allowed to dry in the refrigerator for 1 h.

2.5 Activity assay procedure

Catalase homogeneous activity was determined according to Worthington’s assay procedure [26]. In brief, 100 μL of enzyme solution was added to 2.9 mL of substrate solution (15 mM H₂O₂ in 20 mM buffer, pH 7.00) at a constant temperature of 20°C in a 4.5 mL quartz cuvette. The enzyme kinetics was monitored at 240 nm, at which the variation in absorbance was registered, accepting an extinction coefficient of ε₂₄₀ = 43.6 mol L⁻¹ cm⁻¹. For the determination of the catalytic activity of catalase, 3–5 measurements with the enzyme were carried out. A thermostated spectrophotometer Shimadzu UV–vis 2600 was used.

Electrochemical determination of immobilized catalase activity was performed in two stages. The experiments were carried out in a conventional electrochemical cell with a working volume of 20 mL (in 0.1 M phosphate buffer, pH = 7.0) in a three-electrode setup with a Co₃O₄–g-C₃N₄-modified GCE (developed earlier by our group [25] and proven to be highly sensitive to H₂O₂) as an indicator (working) electrode, a Ag|AgCl, sat. KCl as a reference electrode, and Pt foil as a counter electrode. The measurements were performed at a constant potential of −0.2 V (vs Ag|AgCl) where H₂O₂ electroreduction took place at a significant rate, and no substantial interference from the competing reaction, electrochemical reduction of oxygen [25], was detected.

In the first step, the response of the working electrode, rotating at a constant speed, was recorded as a function of H₂O₂ concentration, which increased gradually through addition of aliquots of H₂O₂ stock solution (0.1 M) to the electrolyte in the cell. The results of measurements were then used to draw the calibration plot for the peroxide catalytic electrode. This procedure was repeated in triplicate in order to ensure sustainability of the electrode response under the experimental conditions. The calibration step will be further denoted in the text as the abiotic stage of the experiment.

In the second biotic stage, the experiment was implemented identically, however, with a glass substrate with immobilized catalase placed on the bottom of the electrochemical cell at ca. 0.8 cm distance from the indicator electrode. The biotic step can be repeated as many times as necessary within the same day.

When examining the immobilized catalase activity in the presence of alcohols, the calibration of the working electrode in the absence of the enzyme was performed with the same concentrations of either methanol, or ethanol, as in the biotic experiments.

3 Results and discussion

Catalase is known to be one of the most efficient biocatalysts with a turnover number being as high as 40 million per second [8]. A widely accepted method for assaying its enzymatic activity in both dissolved and immobilized states is the monitoring of H₂O₂ concentration decay in quiescent solutions, where peroxide is catalytically disproportionated under diffusion control that can potentially result in underestimated activity levels. With this in mind, herein, we discuss an alternative electrochemical approach for the determination of immobilized catalase activity under hydrodynamic conditions.

Figure 1 depicts the hydrodynamic linear sweep voltammograms, recorded on a peroxide-sensitive electrode [25] in the presence of 0.46 mM H₂O₂ at a low scan rate. It can be clearly observed that, over the potential region from −0.3 V to +0.2 V, the electrochemical reduction of H₂O₂ goes under diffusional control. Obviously, the reductive current increases with increasing rotation speed of RDE from 500 to 3,000 rpm. It was found that when the catalytic electrode was rotated at 2,000 rpm, the sensitivity of H₂O₂ determination increased with ca. 25%, as compared with the one estimated in quiescent solutions. Moreover, this sensitivity increase did not cause a considerable growth of the noise level. A further increase of the rotation speed to 3,000 rpm did not result in significant sensitivity improvements; however, the noise levels were much higher, as compared to the results obtained with 2,000 rpm.

Figure 1

Hydrodynamic linear sweep voltammograms recorded in the presence of 0.46 mM H₂O₂ over the catalytic peroxide electrode; rotating speeds, rpm: 500, 832, 1,248, 1,748, 2,332, and 3,000; electrolyte: 0.1 M phosphate buffer, pH = 7.0; temperature: 21 ± 1°C.

During the chronoamperometric experiments aiming to determine the catalytic activity of immobilized catalase performed in quiescent solutions, no observable differences were noted between the electrode current responses recorded in the absence and the presence of catalase up to 4 mM H₂O₂ concentration. It has to be mentioned that the differences in the electrode responses obtained in the abiotic and the biotic stages become obvious only if the working electrode rotates at a considerable rate, thus decreasing the effect of H₂O₂ diffusion through the electrolyte. This finding undoubtedly shows that the diffusion of the enzyme substrate is the rate-limiting step of H₂O₂ decomposition catalyzed by immobilized catalase. All further discussed results were performed at 2,000 rpm rotating speed of the working electrode.

Under the optimized conditions, the response of the catalytic peroxide electrode changed stepwise on addition of H₂O₂ portions, as depicted in Figure 2a. A similar picture is obtained during the second biotic stage, where the experiment was implemented identically, however, with catalase immobilized on a glass substrate placed on the bottom of the electrochemical cell. The dependence of the catalytic peroxide electrode response on the peroxide concentration is presented in Figure 2b. It can be seen that the signal at one and the same peroxide concentration is lower when immobilized catalase was present during the measurements (Figure 2b, blue and red series). The observed effect is a result of decreased H₂O₂ concentration in close proximity to the electrode surface due to the catalytic disproportionation of the latter.

Figure 2

(a) Authentic record of the current variation with time upon addition of H₂O₂ aliquots; operating potential: −0.20 V (vs Ag|AgCl, sat. KCl), PB, pH = 7.0. (b) Dependence of the response of catalytic peroxide electrode on the concentration of H₂O₂ in the electrochemical cell in the absence (black series) and in the presence of 0.25 mg catalase immobilized within a film of Nafion (red series) and cross-linked with glutaric aldehyde (blue series).

The difference between the current response recorded in the abiotic and biotic phases under equivalent other conditions is directly proportional to the rate of H₂O₂ disproportionation, catalyzed by immobilized catalase. This difference plotted versus peroxide concentration represents a non-linear dependence with a hyperbolic trend, which is a characteristic of Michaelis–Menten-type enzyme kinetics. The results obtained during studies on the catalytic behavior of fungal catalase, immobilized in two alternative ways, are compared in Figure 3. The immobilized enzyme through crosslinking with glutaric aldehyde shows a bit higher catalytic activity, while for the catalase entrapped within a layer of Nafion, it was found to be lesser.

Figure 3

Difference between the electrode signal obtained in the abiotic stage and in the biotic phase (ΔI, nA) as a function of H₂O₂ concentration at which it was recorded. Experiments were done with catalase from P. chrysogenum crosslinked with glutaric aldehyde (red series) and entrapped in a Nafion layer (black series) in 0.1 M phosphate buffer, pH = 7.0, at room temperature.

Unfortunately, the substrates with the immobilized enzyme could not be reused, due to an extreme loss of enzymatic activity upon second round of biotic experiments with activity being lost more drastically for the crosslinked enzyme than for the one retained within a polymer membrane. The reason for the observed finding was revealed by SEM–EDX studies, showing that after a single experiment the enzyme amount retained within the polymer film is negligible. In Figure 4 are presented SEM micrographs of immobilized catalase on a conductive support (Al foil) in a polymer layer of Nafion (a), the corresponding EDX spectrum (b), the SEM image of identically immobilized catalase after performing one catalytic cycle (c), and the corresponding EDX spectrum (d). Through comparison of Figure 4a and c, it becomes obvious that the polymer layer is destroyed during the catalytic action of the immobilized catalase, since the smooth-looking surface of the produced heterogeneous biocatalyst turns into a fractured landscape with dendritic formations. The analysis of EDX spectra reveals that the presence of catalase – a Fe-containing enzyme – can be visualized in the spectrum through the Fe peak, presenting ca. 0.7 atomic% in the case of unused immobilized catalase (Figure 4b), and only trace amounts of Fe without the possibility for its quantification can be observed in the case of the already used heterogeneous biocatalyst (Figure 4d). Most probably, the enzyme can escape and leak into the solution during the catalytic experiment. Keeping this in mind, one can speculate that the higher catalytic activity observed when using immobilized catalase via crosslinking is a cumulative result of homogeneous (in solution) and heterogeneous H₂O₂ decomposition, while in the case of catalase immobilized within a polymer film, the observed catalytic activity is mostly due to the heterogeneous peroxide disproportionation.

Figure 4

SEM images of immobilized catalase (a) before and (c) after use in catalytic H₂O₂ disproportionation, and the corresponding EDX spectra showing the presence of enzyme (b) before use and (d) its drastically decreased concentration after use.

All these findings pose the question how reproducible are the measurements if every biotic experiment needs a freshly prepared heterogeneous biocatalyst. In Figure 5 are depicted the experimental results acquired from three consecutive measurements with three identically prepared glass substrates with immobilized catalase. It is obvious that the experimental points practically overlay each other, and the deviation from the mean value does not exceed 2%.

Figure 5

Difference between the electrode signal obtained in the abiotic stage and in the biotic phase (ΔI, nA) as a function of H₂O₂ concentration at which it was recorded for three independent measurements. RSD = 1.96%.

In the late 90s, Magner and Klibanov [27] reported on the unusual catalytic activity of dissolved catalase that mimics that of the peroxidase enzyme; they found that the enzyme is capable of oxidizing alcohols in the presence of H₂O₂ in both aqueous and non-aqueous environments, and the rate of this reaction is manifold higher in organic solvents than in neutral aqueous buffer; however, no additional details about the mechanism of its catalytic action were provided.

By using an already described electrochemical approach, we have been able to monitor the catalytic behavior of immobilized catalase in the presence of 0.5 or 1% low-molecular weight aliphatic alcohols – ethanol and methanol – due to their denaturing effect on the enzyme at higher concentrations, as shown in our recent studies [28].

In Figure 6 are depicted the electrochemically determined H₂O₂ decomposition rates (expressed in nanoamperes) as a function of the concentration, at which it was measured in the presence of 0.5% ethanol or methanol (Figure 6a) and 1.0% of ethanol (Figure 6b). It is obvious that the typical Michaelis hyperbola (shown in Figure 3) turns into a sigmoidal curve when small amounts of alcohols are added to the electrolyte. The sigmoid plot of the dependence is characteristic of the allosteric enzymes capable of binding two distinct substrates at different binding sites, the kinetics of which obeys the Hill equation:

(1) V = V max [ S ] n [ K ] 0.5 n + [ S ] n ,

where V is the rate of the enzyme-catalyzed reaction; V _max is the maximum rate achievable at enzyme saturation with the substrate; K_0.5 is the Hill constant, representing the substrate concentration at which half of the maximum rate is reached; n is the coefficient of Hill, indicative of cooperativity of substrate binding. For n > 1, the cooperativity is positive, i.e., each bond substrate molecule facilitates binding of the next substrate molecule; for n = 1, there is no cooperative binding of substrate, and the Hill equation turns into Michaelis–Menten equation, while for n < 1, there is negative cooperativity, where each bond substrate molecule hampers the binding of the next one.

Figure 6

Difference between the electrode signal obtained in the abiotic stage (I _a, nA) and in the biotic phase (I _b, nA) as a function of H₂O₂ concentration at which it was recorded in the presence of: (a) 0.5% ethanol (black series) or methanol (red series) and (b) 1.0% ethanol. All other conditions are as in Figure 3.

The apparent kinetic constants, calculated from the non-linear regression of the experimental data depicted in Figure 6, are presented in Table 1. It is evident that the apparent kinetic constants are highly dependent on the type of enzyme immobilization – the apparent V _max determined in the absence of alcohol is ca. twice as high for the cross-linked catalase, as compared to the one entrapped within the polymer membrane. However, one can see how detrimental the addition of alcohol to the operating medium is to the enzyme catalytic activity – the crosslinked catalase manifests more than 40 times lower apparent V _max in the presence of 1% ethanol, which is a drastic loss of activity, especially when compared with catalase immobilized via entrapment within a polymeric membrane under equivalent experimental conditions. This finding supports the earlier proposed hypothesis that the cross-linked enzyme can hardly be retained over the glass support and most probably leaks into the solution thus showing mixed activity – heterogeneous and homogeneous.

Table 1

Comparison of the apparent kinetic constants of the immobilized P. chrysogenum catalase immobilized within a Nafion membrane or crosslinked with glutaric aldehyde, as determined electrochemically at H₂O₂ decomposition in the absence and in the presence of 0.5 and 1% ethanol

Kinetic constants	Aqueous buffer, pH = 7.00	0.5% ethanol	1.0% ethanol
*Catalase from P. chrysogenum* immobilized within a film of Nafion^®**
K app , mmol L⁻¹	10.1 ± 0.9	7.2 ± 3.5	2.5 ± 0.5
V max app nA μmol s⁻¹	87.9 ± 5.4	456.6 ± 41.4	7.7 ± 1.5
V max app nA μmol s⁻¹	3.62·10⁻³ ± 2.2·10⁻⁴	18.8·10⁻³ ± 1.7·10⁻³	3.17·10⁻⁵ ± 0.7·10⁻⁵
Hill’s coefficient, n	1.0	4.11 ± 0.27	2.06 ± 0.36
Catalase from P. chrysogenum crosslinked with glutaric aldehyde
K app , mmol L⁻¹	13.1 ± 0.99	8.17 ± 0.5	1.85 ± 0.7
V max app nA μmol s⁻¹	181.2 ± 9.7	243.1 ± 12.5	4.10 ± 0.7
V max app nA μmol s⁻¹	7.46·10⁻³ ± 4.0·10⁻⁴	10.1·10⁻³ ± 0.7·10⁻³	1.70·10⁻⁴ ± 2.9·10⁻⁵
Hill’s coefficient, n	1.0	3.98 ± 0.25	3.36 ± 1.25

Interestingly, the presence of 0.5% ethanol in the solution causes an approximately 5-fold increase in the apparent maximum rate for the catalase immobilized within a polymer film; however, in the case of the crosslinked catalase, this parameter is only slightly higher than the one determined in the absence of ethanol. The apparent Hill’s constant remains close enough to the Michaelis constant determined in the absence of alcohol for both types of enzyme immobilization, while Hill’s coefficient was estimated to be 4, which is not unexpected since the catalase enzyme normally exists as a tetramer.

By increasing the alcohol content to 1.0%, the calculated Hill’s coefficient drops down to 2, the apparent V _max diminishes drastically, and the apparent K_0.5 contracts vastly, which may potentially indicate damage to the enzyme protein shell.

For comparison, another water-soluble peroxide – tert-butyl hydroperoxide – was tested as the catalase substrate in an identical set of experiments; however the enzyme activity toward this particular substrate was calculated to be 597.01 U/mg (ca. one third of the catalase activity toward H₂O₂), and henceforth it will not be subjected to further experiments in the presence of alcohols.

Enzymatic activity of the immobilized catalase was calculated from electrochemical measurements by using the following equation:

(2) A sp = ( ∆ I abiot − ∆ I bio ) ∙ 10 6 s ∙ t V m ,

where A _sp is the specific enzyme activity, U/mg; ΔI _abiot and ΔI _bio, in [A], are the differences between the current recorded at a given concentration with the subtracted background signal for the abiotic and biotic experiments, respectively; s is the sensitivity of the catalytic peroxide electrode (slope of the calibration plot, [A L/mol]); t is the time, [min]; V is the volume of the bioreactor, mL; and m is the amount of the immobilized enzyme [mg].

A comparison of the specific activity of immobilized catalase determined by electrochemical measurements and by spectrophotometry is presented in Table 2. It is evident that the electrochemically determined activity assayed under hydrodynamic conditions is ca. 200 times higher than the one determined under static conditions, most probably resulting from the enhanced mass transport during the electrochemical measurements. Another advantage of the discussed electrochemical approach for the determination of immobilized catalase activity is the opportunity to visualize the alteration of the enzyme kinetic model. The immobilized catalase alters its kinetic pattern from hyperbolic Michaelis type, which is observable in the presence of H₂O₂ only, to sigmoidal Hill’s type describing its catalytic action in the presence of both H₂O₂ and the corresponding hydrogen donor (methanol or ethanol).

Table 2

Specific enzymatic activity of immobilized P. chrysogenum catalase determined electrochemically and spectrophotometrically (n ≥ 3)

% alcohol	A _sp. (el.chem.) in the presence of CH₃OH, μmol min⁻¹ mg⁻¹	A _sp. (spectro) in the presence of CH₃OH, μmol min⁻¹ mg⁻¹	A _sp. (el.chem.) in the presence of C₂H₅OH, μmol min⁻¹ mg⁻¹	A _sp. (spectro) in the presence of C₂H₅OH, μmol min⁻¹ mg⁻¹
0	1672.6 ± 32.7	8.57	1672.6 ± 32.7	8.57
0.5	2033.8 ± 113.8	9.08	1235.5 ± 143.0	7.93
1.0	1076.3 ± 269.1	8.40	537.1 ± 127.5	7.41

It has to be mentioned, however, that the presence of alcohols complicates the measurement process because ethanol and methanol are surfactants that affect both the sensitivity and the precision of the measurements. It can be seen that the higher the percentage of alcohol in operating medium, the bigger the dispersion of the data. Data analysis indicates that the presence of 0.5% methanol enhances the specific enzyme activity with approximately 20% that can be read as a manifestation of peroxidase-like function of catalase, i.e., its ability to oxidize alcohols in the presence of H₂O₂. Such tendency is not observable, however, when ethanol is added in the same percentage, which can be assigned to its more denaturing action on the protein shell of the enzyme.

The observed alterations in immobilized enzyme activity are in agreement with our earlier studies [28], where the activity of “catalase physisorbed on natural fibers” of natural fibers on catalase was assessed spectrophotometrically at much higher H₂O₂ to alcohol ratio.

4 Conclusions

On the basis of the obtained results, the following conclusions can be drawn:

With the help of the discussed electrochemical method, a change of the enzyme catalytic action was observed in the presence of low concentrations of either ethanol or methanol. In the absence of the latter, the immobilized catalase action is characterized by the typical Michaelis–Menten kinetics, while in the presence of either alcohol, the kinetics of its catalytic action obeys Hill’s kinetic model. In addition, the electrochemical monitoring of enzymatic action can be performed in a single experiment, not as time consuming as in the case of spectrophotometrical monitoring. Moreover, the non-enzymatic decomposition of H₂O₂ due to irradiation with UV-light (at 240 nm) is a drawback of the spectrophotometric method that can potentially lead to irreproducibility of the results and incorrect activity assays;
peroxidase-like activity of the immobilized catalase was observed in the presence of methanol in concentrations up to 0.5% (∼130 mM), while the presence of the same concentrations of ethanol suppressed the enzyme activity, most probably due to its stronger denaturing effect on the protein shell of the enzyme.

Summarizing, herein, we demonstrate that an electrochemical method relying on a differential approach – running identical experiments in the absence and presence of an immobilized enzyme – can be successfully employed for quantitative determination of heterogeneous enzymatic activity, provided that the enzyme substrate is electrochemically active and the indicator electrode possesses sufficient sensitivity and selectivity toward it. Another advantage of the discussed method is that enzyme kinetics can be monitored at concentrations within the micromolar range of substrate concentrations, which is not possible to be done spectrophotometrically.

Acknowledgements

Electrochemical studies were performed thanks to the research infrastructure of the Center for Competence “Personalized Innovative Medicine, PERIMED (grant BG05M2OP001-1.002-0005-C01)”.

Funding information: This research was funded by the Bulgarian National Science Fund (grant KP-06-N39/8).
Author contributions: Angel Peshkov – investigation, methodology, data curation, writing – original draft; Mariya Pimpilova – methodology, data curation, writing – original draft; Merin Shukri – investigation, data curation; Ilia Iliev – resources, supervision, writing – review and editing. Nina Dimcheva – conceptualization, funding acquisition, project administration, formal analysis, supervision, writing – review and editing. All authors have read and agreed to the published version of the manuscript.
Conflict of interest: The authors state that there are no conflicts to declare.
Ethical approval: The conducted research is not related to either human or animals use.
Data availability statement: All data can be provided by the corresponding author (N.D.) upon reasonable request.

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Received: 2024-10-29

Revised: 2025-02-05

Accepted: 2025-03-07

Published Online: 2025-03-29

This work is licensed under the Creative Commons Attribution 4.0 International License.

Artikel in diesem Heft

https://doi.org/10.1515/chem-2025-0143

Schlagwörter für diesen Artikel

immobilized catalase; enzyme kinetics; alcohols; peroxidase-like activity; electrochemical method

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