Characterization of pyranose oxidase variants for bioelectrocatalytic applications

Abstract

Pyranose oxidase (POx) catalyzes the oxidation of d-glucose to 2-ketoglucose with concurrent reduction of oxygen to H₂O₂. POx from Trametes ochracea (ToPOx) is known to react with alternative electron acceptors including 1,4-benzoquinone (1,4-BQ), 2,6-dichlorophenol indophenol (DCPIP), and the ferrocenium ion. In this study, enzyme variants with improved electron acceptor turnover and reduced oxygen turnover were characterized as potential anode biocatalysts. Pre-steady-state kinetics of the oxidative half-reaction of ToPOx variants T166R, Q448H, L545C, and L547R with these alternative electron acceptors were evaluated using stopped-flow spectrophotometry. Higher kinetic constants were observed as compared to the wild-type ToPOx for some of the variants. Subsequently, the variants were immobilized on glassy carbon electrodes. Cyclic voltammetry measurements were performed to measure the electrochemical responses of these variants with glucose as substrate in the presence of 1,4-BQ, DCPIP, or ferrocene methanol as redox mediators. High catalytic efficiencies (I_max^app/K_M^app) compared to the wild-type POx proved the potential of these variants for future bioelectrocatalytic applications, in biosensors or biofuel cells. Among the variants, L545C showed the most desirable properties as determined kinetically and electrochemically.

Highlights

• •Pyranose oxidase (POx) shows attractive features for bioelectrocatalysis.
• •Trametes ochracea POx variant L545C is most promising for these applications.
• •Rapid kinetics experiments give good predictions for performance on an electrode.

1. Introduction

Bioelectrocatalysis as a special section of biocatalysis has been increasingly exploited and studied in recent years in order to develop new or improved bioelectrochemical devices, including biofuel cells (BFCs) or biosensors. In accordance to biocatalysis, where enzymes (or cells) are used as biocatalysts for the production of chemicals, bioelectrocatalysis is employing enzymes – and here predominantly oxidoreductases – for the generation of electricity or analytical signals. Bioelectrocatalysis is thus a key element in these applications, as electrons involved in an enzymatic reaction are collected at an electrode surface. Especially BFCs are slowly coming of age as next generation power sources as is evident from significant improvements and scientific progress over the last years [1]. In contrast to conventional fuel cells, BFCs use more diverse fuels and offer the opportunity of using more complex molecules as energy sources at modest and biocompatible conditions. Anodic bioelectrocatalysis is the study on enzymes responsible for the oxidation of fuels in BFCs, where electrical energy is generated from chemical energy upon the oxidation of the fuel, typically a sugar or an alcohol [2]. The most frequently used enzyme for these anodic applications in BFCs has been glucose oxidase (GOx), yet lately a number of alternative oxidoreductases emerged that show better or improved properties compared to GOx, such as a wider substrate (fuel) spectrum, promiscuous activities on several substrate molecules, reduced/negligible activity with oxygen, or direct electron transfer [2,3]. One such alternative enzyme for BFC applications is pyranose oxidase (POx) (pyranose:oxygen 2-oxidoreductase; EC 1.1.3.10), which catalyzes the oxidation of several aldopyranoses at their C2 position to produce an α-ketopyranose with simultaneous reduction of oxygen to H₂O₂. Although d-glucose is the preferred substrate, POx can also catalyze the oxidation of other carbohydrates including d--galactose, l--sorbose, d--xylose, and d--glucono-1,5-lactone [4]. POx has been isolated and purified from several ascomycetes like Aspergillus nidulans and Aspergillus oryzae [5], basidiomycetes including Phanerochaete chrysosporium [6], Phlebiopsis gigantea [7] and Trametes ochracea [8] (formerly known as Trametes multicolor), and lately also from bacteria such as Arthrobacter siccitolerans [9] and Kitasatospora aureofaciens [10].

The reaction catalyzed by POx can be divided into two half-reactions. In the reductive half-reaction, two electrons, as hydride equivalent, are donated by the sugar substrate to POx(FAD) yielding POx(FADH₂) and the oxidized sugar. In the ensuing oxidative half-reaction, two electrons are transferred to oxygen to produce H₂O₂ and regenerate POx(FAD) [11]. Alternatively, POx can also react with other electron acceptors; in fact some of these show catalytic behaviour superior to that with oxygen [12].

The currently best-studied POx is from Trametes ochracea (ToPOx), which has been well characterized from a kinetic and mechanistic [[13], [14], [15]] as well as from a structural [[16], [17], [18]] point of view. In order to tailor ToPOx for applications in BFCs, we engineered this enzyme to decrease oxygen reactivity [19]. The aim of this previous engineering programme based on structural insight was to decrease the activity of ToPOx with oxygen (thus reducing or avoiding the formation of hydrogen peroxide) while maintaining (or even improving) the activity with the alternative electron acceptors 1,4-BQ, DCPIP and the ferrocenium ion. Eleven residues of ToPOx were selected based on their vicinity to the flavin cofactor and targeted by site-saturation mutagenesis, thereby creating all possible variants at these positions. ToPOx variants T166R, Q448H, L545C, L547R and N593C were identified in microtiter plate-based screening assays, in which the activity with oxygen and the alternative electron acceptors was measured, and preliminarily characterized by determining their apparent steady-state kinetic constants for different electron acceptors [19]. In general, selected variants are characterized by significantly reduced activities with oxygen (relative activities ranging from 0.06 to 39% compared to wild-type ToPOx as judged from the catalytic efficiencies (k_cat/K_M), while activity with the alternative electron acceptors was in some cases reduced, maintained or even considerably increased [19]. Furthermore, we employed ToPOx N593C, which showed the most pronounced reduction in oxygen reactivity, for detailed electrochemical characterization [20].

The aim of this current study was the detailed biochemical and bioelectrochemical characterization of the ToPOx variants T166R, Q448H, L545C, and L547R as well as their comparison with wild-type (WT) POx pertaining to the reactivity with alternative electron acceptors. Fig. 1 shows the position of the mutated amino acid residues in the close vicinity of the isoalloxazine ring of FAD. Overall, we were interested whether these variants show promise for applications in bioelectrocatalysis and BFCs.

Fig. 1.

Alignment of the active-site architecture of Trametes ochracea pyranose oxidase wild-type (PDB 1TT0) as well as of the modeled structures of variants T166R (red), Q448H (green), L545C (blue), and L547R (orange). The isoalloxazine moiety of FAD and the sugar substrate-mimicking acetate in the active site are shown in black and cyan, respectively. Original residues are shown in yellow. For clarity not all residues are shown and colored. The image was generated using PyMOL. Structures of variants were generated using Molecular Operating Environment (MOE, Chemical Computing Group Inc., Montreal, Quebec Canada).

2. Materials and method

2.1. Chemicals

Chitosan, ferrocene methanol (MeOHFc), horseradish peroxidase, and phenyl methyl sulfonyl fluoride (PMSF) were purchased from Sigma. 1,4-Benzoquinone (1,4-BQ) and ferrocenium hexafluorophosphate (FcPF₆) were obtained from Aldrich. d-glucose, potassium dihydrogen phosphate, potassium phosphate, ammonium sulfate, and NaCl were purchased from Roth. Imidazole was obtained from AppliChem; 2,6-dichlorophenol indophenol (DCPIP) from Fluka; ABTS [2,2′-azinobis(3-ethylbenzthiazolinesulfonic acid)] from Amresco; and glutaraldehyde (GA) from Merck.

2.2. Gene expression and recombinant protein purification

ToPOx (wild-type and variants) was recombinantly produced in E. coli as described previously [19]. In brief, ToPOx was expressed in E. coli BL21*DE3 grown in 1 L TB_amp medium (separated in four flasks each containing 250 mL of medium) at 37 °C and 160 rpm. When OD₆₀₀ of 0.5 was reached, the expression was induced by adding a lactose solution (150 g L⁻¹) to give a final concentration of 5 g L⁻¹ medium. Incubation was continued for 20 h at 25 °C and 160 rpm. Centrifugation (5000 rpm, 20 min, 4 °C, Beckman Coulter Avanti J26 XP) was done to collect the cell pellets. The pellets were suspended in buffer A (50 mM phosphate buffer, 50 mM NaCl, 50 mM imidazole, pH 6.5), to which 10 μL of PMSF (10 mg mL⁻¹) was added for every 10 mL of cell pellet solution. A French press (1000 bar) was used to disrupt the cells. The crude extract was collected after centrifugation (25,000 rpm, 30 min, 4 °C, Beckman Coulter Avanti J26 XP). This was then loaded onto IMAC Ni-charged resin (5 mL, HiTrap IMAC HP, GE Healthcare Life Sciences) followed by washing with 3 column volumes of buffer A to remove unbound proteins. The enzyme was eluted with a linear gradient of buffer B (50 mM phosphate buffer, 500 mM NaCl, 500 mM imidazole, pH 6.5). Active fractions (as measured by the ABTS assay) showing clear absorbance at 280 and 460 nm were collected and concentrated via centrifugation (4000 rpm, 30 min, 4 °C, Eppendorf 5810R) with an Amicon Ultra Centrifugal Filter Device of 100-kDa cut-off (Millipore).

2.3. Protein analysis

Protein concentrations were determined using Bradford's method with the Bio-Rad Protein assay Kit containing bovine serum albumin as standard. SDS-PAGE was performed using Bio-Rad Mini-PROTEAN TGX stain-free gels.

2.4. Enzyme activity assay

The peroxidase-coupled assay based on ABTS was used in the standard measurement of POx activity. Assay mixtures (980 μL total volume) were prepared by combining 1 μmole ABTS, 2 U of horseradish peroxidase, 100 μmole d-glucose in 50 mM potassium phosphate buffer (KPP) buffer (pH 6.5). The assay mixture was pre-warmed at 30 °C, and the reaction started by adding 20 μL of enzyme solution to the assay mixture. The activity of POx was determined using a spectrophotometer at 400 nm by measuring the amount of H₂O₂ formed for 3 min (ε_ABTS = 43.2 mM⁻¹ cm⁻¹). One unit of POx activity is defined as the amount of POx needed for the oxidation of 2 μmol of ABTS [8].

2.5. Rapid kinetics experiments

Rapid kinetics experiments were performed using an Applied Photophysics SX-20 stopped-flow spectrophotometer (Applied Photophysics, Leatherhead, UK) in single-mixing mode. The optical path length of the observation cell was 1 cm. Before measurements, the stopped-flow equipment was rinsed with anaerobic buffer. Enzyme solutions and substrates at various concentrations were placed in a tonometer and were made anaerobic by alternatively introducing vacuum and oxygen-free nitrogen for 15 min [14]. All measurements were carried out in 50 mM KPP buffer (pH 6.5) at 25 °C. The enzyme solutions (40 μM) were reduced with a solution of 4 mM d-glucose in KPP in a tonometer. The reduced enzyme was loaded onto the stopped-flow photometer and allowed to react with DCPIP (0.030–2.00 mM), 1,4-BQ (0.005–0.500 mM) and Fc⁺ (0.050–2.00 mM). Reactions were monitored at various wavelengths of 200–700 nm for 0.00126 to 60 s using a photodiode-array detector. Apparent rate constants were analyzed using the Pro-Data SX software (Applied Photophysics). The resulting plots of absorbance vs. time were fit to a single exponential equation corrected for offset (Eq. (1)).

$$y=A^{-k_{\mathit{obs}}t}+c$$ (1)

Maximum observed rate constants (k_{obs, max}) were calculated by non-linear least-square regression, fitting the data of observed rate constants against concentrations of redox mediators to the One Site Saturation equation in SigmaPlot (Systat Software Inc., San Jose, CA).

2.6. Preparation of the enzyme-modified electrodes

Glassy carbon electrodes (GCE, Bioanalytical Systems, West Lafayette, IN, USA; 3.0 mm diameter) were polished with alumina slurries (0.3 and 0.05 mm, Micro Polish) on microcloth (Buehler), and then sonicated in deionized water for 3 min to remove particles. Enzyme mixtures were prepared by mixing 100 μL of the enzyme (8 mg mL⁻¹, in 50 mM sodium citrate buffer, pH 5.0), 100 μL of chitosan (2 mg mL⁻¹, pH 5.0) and 10 μL of GA (5%). The mixture was vortexed for 1 min, this enzyme solution (5 μL) was dropped onto the polished electrode surface, and allowed to dry overnight at 4 °C and constant humidity. The electrodes were then stored at room temperature for complete drying.

2.7. Electrochemical measurements

Cyclic voltammetry (CV) measurements were carried out in 50 mM potassium phosphate buffer (pH 6.5) containing 100 mM KCl (PBS). Solutions were purged with nitrogen for 15 min before measurements to avoid any possible interference by residual oxygen. The electrode responses were measured against a commercial Ag∣AgCl∣KCl (sat., aqu.) reference electrode and a platinum wire electrode as the counter electrode using a three-electrode potentiostat (Metrohm Autolab PGSTAT204). CV was performed at the scan rate of 10 mV s⁻¹ from −100 mV to +400 mV for 1,4-BQ; −50 to +500 mV for MeOHFc; and −150 to +300 mV for DCPIP solutions with or without glucose [21,22]. Catalytic currents obtained with redox mediators were taken from CVs at the following potentials: 1,4-BQ (250 mV), MeOHFc (300 mV), and DCPIP (125 mV) vs. Ag∣AgCl, which are below the anodic current maximum, but therefore also not mass-transfer limited. Diffusion constants for the three mediator 1,4-BQ, MeOHFc and DCPIP are 3.26 × 10⁻⁹ m² s⁻¹ (in aqueous medium), 3.1× 10⁻⁹ m² s⁻¹ (in propylene carbonate medium) and 0.77 × 10⁻⁹ m² s⁻¹ (in aqueous medium), respectively [23,24].

3. Results

3.1. Generation of ToPOx variants

Both wild-type ToPOx and the ToPOx variants T166R, Q448H, L545C and L547R were recombinantly produced in E. coli, and the His-tagged proteins were purified by Ni-immobilized metal ion affinity chromatography (IMAC) after cell disruption. SDS-PAGE analysis (Fig. 2) confirmed the purity and identity of the bright yellow fractions collected from IMAC. All purified fractions (WT and variants) showed a dominant and distinct protein band at around 65 kDa, corresponding to the molecular mass of the single ToPOx subunit.

Fig. 2.

SDS-PAGE of wild-type POx and different variants. Lane 1, molecular mass standard Precision Plus Protein Stained (Bio-Rad); Lane 2, WT ToPOx; Lane 3, T166R; Lane 4, Q448H; Lane 5, L545C; and Lane 6, L547R.

3.2. Kinetics of the oxidative half-reaction of ToPOx variants

The oxidation of reduced POx(FADH₂) was observed by following the increase in absorbance at 459 nm (Fig. 3). Anaerobic solutions of the reduced enzyme were mixed with solutions containing various concentrations of the electron acceptors 1,4-BQ, Fc⁺ and DCPIP using a stopped-flow spectrophotometer. The pre-steady state measurements started at 0.00126 s, and the increase in absorbance at 459 nm was followed until 0.2 s for all the electron acceptor substrates tested. The observed rate constants (k_obs) for this phase of the reaction were calculated by exponential fitting to Eq. (1) for each concentration of electron acceptor applied in the experiment. The hyperbolic dependence of k_obs on the concentration of electron acceptors suggests that at least one more step preceding flavin oxidation exists [11]. Overall, the reaction phase involves the electron transfer from reduced flavin [POx(FADH₂)] to the electron acceptor (redox mediator RM in its oxidized form). RM first forms a complex with the flavin (of unknown structure and depending on the electron acceptor), which is then oxidized with the simultaneous reduction of RM to RM_red. This is immediately followed by the release of RM_red to yield oxidized POx(FAD). We calculated the maximum observed rate constants (k_{obs, max}), which characterise the overall reaction of the reduced enzyme with the mediators (Schemes 1 and 2), for the different mediators and variants. Results are summarized in Table 1.

Fig. 3.

Kinetic traces from the absorbance at 459 nm of the oxidation of reduced POx (WT and variants) by (A) 0.05 mM 1,4-BQ, (B) 0.05 mM Fc⁺, and (C) 0.03 mM DCPIP.

Table 1.

Maximum observed rate constants (k_obs,max) of the oxidative half-reaction of wild-type and mutant To POx for the electron acceptors 1,4-benzoquinone (1,4-BQ), 2,6-dichlorophenolindophenol (DCPIP) and ferrocenium ion (Fc⁺), the concentrations of which were varied in a series of experiments. Reactions were performed at 25 °C.

Enzyme	kobs,max ± SD∗
	1,4-BQ	Fc+	DCPIP
WT	71.1 ± 6.9	50.7 ± 9.1	65.5 ± 8.7
T166R	124 ± 6	135 ± 12	23.5 ± 4.8
Q448H	36.6 ± 2.1	80.7 ± 18.4	32.8 ± 5.8
L545C	426 ± 9	425 ± 113	62.9 ± 8.5
L547R	136 ± 21	57.8 ± 8.1	20.9 ± 3.6

SD = standard deviation calculated from three independent experiments.

Scheme 1: (two-electron acceptor)

$$\mathit{POx}\left({\mathit{FADH}}_2\right)+\mathit{RM}\stackrel{k_{\mathit{obs}\mathit{max}}}{\to }\mathit{POx}\left(\mathit{FAD}\right)+{\mathit{RM}}_{\mathit{red}}$$

Scheme 2: (one-electron acceptor)

$$\mathit{POx}\left({\mathit{FADH}}_2\right)+2\mathit{RM}\stackrel{k_{\mathit{obs}\mathit{max}}}{\to }\mathit{POx}\left(\mathit{FAD}\right)+2{\mathit{RM}}_{\mathit{red}}$$

In general, all variants reacted faster with 1,4-BQ and Fc⁺ when compared to the wild-type enzyme (with the exception of Q448H and 1,4-BQ). Overall, L545C shows the highest maximum observed rate constants with these two mediators as compared to WT, with 6.0- and 8.4-fold increases in k_{obs, max} for 1,4-BQ and Fc⁺, respectively, relative toWT.

3.3. Electrochemical characterization of POx immobilized on electrodes

First, cyclic voltammograms of POx-modified glassy carbon electrodes (GCE) in the presence of different redox mediators and in the absence of glucose were obtained. The applied potential was varied at the low scan rate of 10 mV s⁻¹ to ensure steady-state behaviour. The modulation of the potential governs the electron transfer between the working electrode and the mediator species in the solution, and causes the oxidation or reduction of the mediators in vicinity of the electrode as indicated by the presence of anodic and cathodic peaks in the cyclic voltammograms. Fig. 4A shows the voltammograms obtained for WT-GCE in the presence of different redox mediators serving as co-substrates and electron acceptors of POx. The scanned potential window was varied for each redox mediator depending on its redox potential.

Fig. 4.

Recorded voltammograms (scan rate: 10 mV s⁻¹) of (A) WT-GCE in the presence of various mediators, and (B—F) POx-modified GCE in the presence of varying concentrations of 1,4-benzoquinone in nitrogen-saturated solutions of 0.1 M potassium phosphate buffer at pH 6.5.

The CV of WT-modified GCE in the presence of 1.0 mM 1,4-BQ (Fig. 4A) shows the two-electron/two proton redox reaction of 1,4-BQ/hydroquinone (HQ) (Eq. (2)). The anodic (E_pa) and cathodic peak potentials (E_pc) are at 210 and 5 mV, respectively, and the peak separation (ΔE_p) is 205 mV. The oxidative current starts at approximately 150 mV, pointing towards the redox reaction of 1,4-BQ/HQ and not the oxidation of FAD. FAD oxidation in GOx occurred at about −50 to −250 mV depending on the pH of the solution [25], and the redox potential of free FAD in solution is at −207 mV (pH 7.0) [26]. The large peak-to-peak separation observed in the buffered solution near neutral pH indicates that the reaction is thermodynamically very favourable, but the electron transfer is not inherently fast, as is generally the case with quinones in aqueous solutions [27]. The recorded CV of WT-modified GCE in the presence of 1.0 mM MeOHFc (Fig. 4A) shows the one-electron/no-proton redox reaction of MeOHFc/MeOHFc⁺ (Eq. (3)) with E_pa at 285 mV and E_pc at 212 mV. The peak separation ΔE_p is 73 mV. The anodic and cathodic peaks occurred at a potential typical for the electron transfer involving the redox reaction of MeOHFc/MeOHFc⁺ [28,29]. For the CV profile of WT-GCE in the presence of 1.0 mM DCPIP (Fig. 4A), the anodic and cathodic peaks represent the two-electron/two-proton redox reaction of DCPIP/DCPIPH₂ (Eq. (4)). The E_pa and E_pc are at 128 and 61 mV, respectively, and the peaks are separated by 67 mV. The oxidation starts at about 50 mV corresponding to oxidation of reduced DCPIP at the base of the electrode.

$$1,4-\mathit{BQ}+2e^{-}+2H^{+}\rightleftharpoons \mathit{HQ}$$ (2)

$$\mathit{MeOHFc}\rightleftharpoons \mathit{MeOH}{\mathit{Fc}}^{+}+e^{-}$$ (3)

$$\mathit{DCPIP}+2H^{+}+2e^{-}\rightleftharpoons \mathit{DCPIP}{H}_2$$ (4)

Upon addition of 50 mM glucose to solutions containing various concentrations of 1,4-BQ (0.01–2.0 mM), the CV profiles showed the typical sigmoidal catalytic curves corresponding to glucose oxidation (Fig. 4B to F) [30]. The dramatic increase in the oxidation peak with increasing 1,4-BQ concentrations is proportional to the efficiency of glucose oxidation. The cathodic current completely (or almost completely) disappeared for the WT- and variant-modified GCE. This disappearance of the cathodic peak indicates that 1,4-BQ reacts more efficiently with reduced POx (Eq. (6)) compared to the reduction of 1,4-BQ at the electrode. This implies that WT, Q448H, and L545C show higher catalytic activity than the other POx variants with 1,4-BQ when immobilized on the electrode since the cathodic peak disappeared for these three POx-modified electrodes, whereas the cathodic peak reappeared at higher 1,4-BQ concentrations for T166R and L547R. These cathodic peaks also increased with higher concentrations of 1,4-BQ, suggesting that the POx activity present is not sufficient for the complete reduction of 1,4-BQ anymore and that a certain fraction of 1,4-BQ is reduced by the electrode.

Fig. 5 shows the bioelectrocatalytic response of ToPOx-modified GCEs with a constant amount of glucose and varying concentrations of MeOHFc. Again, the CV profile changed when glucose was present together with the mediator forming a well-defined catalytic sigmoidal curve, which is similar to the electrocatalytic response of glucose oxidation by glucose oxidase in the presence of MeOHFc [31]. Addition of glucose resulted in a significant increase of the oxidation peak and disappearance of the reduction current. This again shows the very good reactivity of ToPOx with the mediator MeOHFc, as it is more efficiently reduced by POx(FADH₂) (Eq. (7)) than by electroreduction.

Fig. 5.

Recorded voltammograms (scan rate: 10 mV s⁻¹) POx-modified GCE with varying concentrations of ferrocene methanol in nitrogen-saturated solutions of 0.1 M potassium phosphate buffer at pH 6.5.

Comparable changes were also observed in the CV profiles of POx-modified GCE upon addition of glucose to DCPIP solutions (Fig. 6A–E). Reduced POx (both WT and variants) efficiently transferred electrons to DCPIP (Eq. (8)), which diminished or completely abolished the cathodic peak corresponding to DCPIP reduction at the electrode (Eq. (4)). We noticed that DCPIP strongly adsorbs onto the POx-modified electrode surface after extensive and repeated use, and cannot be removed by rinsing or immersion in a buffer solution between measurements. This adsorption of DCPIP onto the electrode was verified by CVs measured with these electrodes, previously used with DCPIP, in the presence of glucose but absence of DCPIP added to the buffer. These electrodes gave CVs with similar response curves as those with soluble DCPIP, albeit with much lower currents (Fig. 6F). To limit the electrodeposition of DCPIP on the electrode, which would affect mass-transfer and therefore current response, the electrodes were frequently exchanged especially in CV measurements with high DCPIP concentrations [32].

$$\mathit{POx}\left(\mathit{FAD}\right)+\mathit{glucose}\to 2-\mathit{KG}+\mathit{POx}\left(\mathit{FAD}{H}_2\right)$$ (5)

$$\mathit{POx}\left(\mathit{FAD}{H}_2\right)+1,4-\mathit{BQ}\stackrel{k}{\to }\mathit{POx}\left(\mathit{FAD}\right)+\mathit{HQ}$$ (6)

$$\mathit{POx}\left(\mathit{FAD}{H}_2\right)+2\mathit{MeOH}{\mathit{Fc}}^{+}\stackrel{k}{\to }\mathit{POx}\left(\mathit{FAD}\right)+2\mathit{MeOHFc}$$ (7)

$$\mathit{POx}\left(\mathit{FAD}{H}_2\right)+\mathit{DCPIP}\stackrel{k}{\to }\mathit{POx}\left(\mathit{FAD}\right)+\mathit{DCPIP}{H}_2$$ (8)

Fig. 6.

Recorded voltammograms (scan rate: 10 mV s⁻¹) of POx-modified GCE with varying concentrations of 2,6-dichlorophenol indophenol in nitrogen-saturated solutions of 0.1 M potassium phosphate buffer at pH 6.5.

3.4. Kinetic characterization of enzyme-modified electrodes

The anodic current generated by the oxidation of glucose in the presence of a mediator is related to the catalytic efficiency or turnover number of the enzyme adsorbed onto the electrode [33]. This can be used for the kinetic characterization of the electrode-bound enzymes and the determination of apparent kinetic constants, by plotting the catalytic current for the oxidation of glucose versus the different concentrations of mediator employed [30,34]. In these experiments, the concentration of glucose was constant and present in excess and thus saturating concentrations (at least a 25-fold higher concentration in relation to the redox mediator). Since glucose-saturating conditions can, however, not be guaranteed throughout the experiments, especially within the chitosan layer, data here are reported as apparent kinetic constants (I_max^app, K_M^app) in accordance with the reporting of biochemical kinetic data (k_cat, K_M) obtained in steady-state experiments. The highest I_max^app values were obtained with WT ToPOx as well as with variants Q448H and L545C when using 1,4-BQ as mediator. Furthermore, all of the variant-modified GCE showed lower apparent K_M^app values with 1,4-BQ compared to the WT-GCE. In addition, they also showed higher catalytic current efficiencies I_max^app/K_M^app (Table 2). T166R and L547R showed the most significant decrease of K_M compared to WT ToPOx, which resulted in an increased catalytic efficiency (4.3- and 2.6-fold higher relative to WT ToPOx, respectively). However, these two variants also showed lower maximum current values (I_max^app) than the WT. This should lead to the generation of lower power outputs when applying T166R and L547R together with 1,4-BQ in BFC applications. Variants Q448H and L545C show K_M values comparable to the WT enzyme and modestly higher catalytic efficiencies compared to the WT (1.5- and 1.4-fold higher, respectively).

Table 2.

Steady-state kinetic constants of wild-type and mutant ToPOx with 1,4-benzoquinone as the electron acceptor and 50 mM d-glucose as the electron donor. The I_max^app/K_M^app value is reported relative to the value calculated for the wild-type enzyme, which is given as 100%.

Enzyme	KMapp ± SD (mM)	Imaxapp ± SD (μA)	Imaxapp/KMapp (%)	KM∗ (mM)	KMapp/KM∗
WT	1.332 ± 0.456	12.5 ± 2.1	100	0.24 ± 0.03a	5.6
T166R	0.170 ± 0.046	6.8 ± 2.7	429	0.14 ± 0.02b	1.2
Q448H	1.196 ± 0.241	16.3 ± 4.9	145	0.057 ± 0.007	21.0
L545C	0.954 ± 0.114	12.2 ± 0.9	136	0.042 ± 0.006b	22.7
L547R	0.371 ± 0.017	9.2 ± 2.6	264	0.018 ± 0.02b	20.6

SD = standard deviation calculated from three replicated experiments.

^aData are from [42].

^bData from [19].

When using DCPIP as an electron acceptor, L545C showed a low K_M^app and the highest I_max^app values for this mediator, which together resulted in an approx. 2.9-fold higher catalytic efficiency compared to the WT (Table 3). T166R and Q448H also showed better K_M^app values while I_max^app was comparable to the WT enzyme. This resulted in approx. 1.8- and 2-fold higher catalytic efficiency for T166R and Q448H, respectively. L547R showed both a high I_max^app and a high K_M^app (about 1.7-fold higher than WT), which gave an overall performance comparable to WT (similar catalytic efficiency).

Table 3.

Steady-state kinetic constants of wild-type and mutant ToPOx with 2,6-dichlorophenol indophenol as the electron acceptor and 50 mM d-glucose as the electron donor. The I_max^app/K_M^app value is reported relative to the value calculated for the wild-type enzyme, which is given as 100%.

Enzyme	KMapp ± SD (mM)	Imaxapp ± SD (μA)	Imaxapp/KMapp (%)	KM∗ (mM)	KMapp/KM∗
WT	1.020 ± 0.277	31.3 ± 3.0	100	1.9 ± 0.1a	0.5
T166R	0.546 ± 0.073	29.7 ± 1.4	177	2.1 ± 0.5	0.3
Q448H	0.641 ± 0.096	39.3 ± 2.5	200	0.24 ± 0.03	2.7
L545C	0.561 ± 0.112	49.5 ± 6.4	287	0.72 ± 0.07	0.8
L547R	1.699 ± 0.030	49.4 ± 6.1	95	1.7 ± 0.4	1.0

SD = standard deviation calculated from three replicated experiments.

^aData from [19].

All the ToPOx variants except Q448H showed increased K_M^app values with MeOHFc (1e⁻/non-H⁺ acceptor) (Table 4). Despite this unfavourable K_M^app value, L545C showed both the highest maximum current I_max^app among the four variants (a 19-fold increase compared to WT) as well as the highest catalytic current efficiency (4.3-fold increase than WT) when using MeOHFc. This could be very attractive for BFC applications since higher power outputs can be obtained by a high current output, and ferrocenes are frequently used as mediators in bioelectrochemical applications.

Table 4.

Steady-state kinetic constants of wild-type and mutant ToPOx with ferrocene methanol as the electron acceptor and 50 mM d-glucose as the electron donor. The I_max^app/K_M^app value is reported relative to the value calculated for the wild-type enzyme, which is given as 100%.

Enzyme	KMapp ± SD (mM)	Imaxapp ± SD (μA)	Imaxapp/KMapp (%)
WT	0.288 ± 0.014	12.1 ± 0.9	100
T166R	0.933 ± 0.086	67.0 ± 17.3	171
Q448H	0.144 ± 0.023	2.7 ± 0.7	45
L545C	1.290 ± 0.076	231 ± 10	427
L547R	0.731 ± 0.040	41.4 ± 6.8	135

SD = standard deviation calculated from three replicated experiments.

Generally, the apparent Michaelis constants of enzymes on electrodes are higher than those in solutions [19]. The ratio of the Michaelis constants of electrode-immobilized POx (K_M^app) and free soluble enzyme (K_M^⁎) is given by (K_M^app/K_M^∗) in Table 2, Table 3. Interestingly, we did not observe a general increase in K_M for the electrode-immobilized enzyme, which could be assigned to a general mass-transfer restriction imposed by the chitosan layer. An increase was observed for both WT and the ToPOx variants when using 1,4-BQ, albeit the K_M^app/K_M^∗ ratio varied considerable for different variants, with this ratio going up to 22.7 for L545C. This is in contrast to the electron acceptor substrate DCPIP, where the Michaelis constant is always comparable for the electrode-bound and the free enzyme (K_M^app/K_M^∗ ranging from 0.3 to 2.7). Since we used different ferrocenes for the biochemical and bioelectrochemical characterization (ferrocenium hexafluorophosphate and ferrocene methanol, respectively) the K_M^app/K_M^∗ ratio cannot be given for this electron acceptor.

4. Discussion

Biological fuel cells, and more specifically enzyme fuel cells based on oxidoreductases, hold promise as small, lightweight and sustainable power sources that use simple renewable fuels for possible applications in consumer electronics or implantable electronics. Their operational advantages over alternative technologies for generating power from organic fuels are the high conversion efficiency as well as their operation under ambient conditions. Yet, some major limitations still hamper a wider successful introduction into various products, and these challenges include relatively low power densities, poor operational stability and limited voltage output [1]. Power densities of BFC may be enhanced by improving the kinetic properties of an enzyme, which can be achieved by increasing the rates of electron transfer between the oxidoreductase and the electrode (higher k_cat) or by improving the catalytic efficiency of an enzyme (higher k_cat/K_M). One factor contributing to increased stability is the avoidance of hydrogen peroxide formation, which can be achieved by employing dehydrogenases rather than oxidases as anodic biocatalysts, or by engineering oxidases so that their oxygen reactivity is tuned down [1]. Alternatively, enzymes can be engineered to increase their (thermo)stability by using various approaches of enzyme evolution [35,36].

We recently engineered ToPOx by mutating amino acid residues around the active site and the isoalloxazine ring (Fig. 1), and screened for variants showing reduced activity with oxygen while activity with selected alternative electron acceptors representing different groups of mediators (1,4-BQ, Fc⁺, DCPIP) should be retained. This resulted in the identification of variants T166R, Q448H, L545C and L547R [19]. As judged from steady-state kinetic analysis and apparent steady-state kinetic constants, these variants showed significantly reduced catalytic efficiencies with oxygen (relative k_cat/K_M values of <1–39% compared to WT), while the activity with the alternative electron acceptors were well retained and in some instances even increased in relation to the WT.

In order to study the reaction of ToPOx with these different electron acceptors in more detail we performed rapid kinetic experiments, in which the reduced enzyme reacted with 1,4-BQ, DCPIP and Fc⁺ of varying concentrations, and calculated the maximum observed rate constants k_{obs, max}, which characterise the overall reaction of the reduced enzyme with the mediators, for this oxidative half-reaction. Some of the ToPOx variants showed significantly higher k_{obs, max} values for the electron acceptors compared to the wild-type, with L545C showing the most pronounced improvement of k_{obs, max} (6.0 and 8.4 -fold improved values for 1,4-BQ and Fc⁺, respectively, compared to the WT). An improvement of the rate constant was not always consistent for the different variants, e.g., k_{obs, max} was increased relative to the WT for Q448H when using Fc⁺, while it was decreased for both 1,4-BQ and DCPIP.

Immobilisation of POx on glassy carbon electrodes involved cross-linking with glutaraldehyde and chitosan. This may affect the catalytic properties of an enzyme significantly, as for example it may hamper access of a substrate to the active site or orient an enzyme in an unfavourable way. Therefore we also determined the apparent kinetic constants of the electrode-immobilized enzymes for the electron acceptors/redox mediators 1,4-BQ, DCPIP and MeOHFc. In accordance with the rapid kinetic data, some of the variant showed kinetic properties that are more favourable than those of the WT, and again L545C exhibited the most significant improvements, in particular when using DCPIP and MeOHFc as mediator substrate. Here both the maximum current I_max^app as well as the catalytic current efficiency I_max^app/K_M^app are improved considerably. In addition, T166R showed improved I_max^app/K_M^app values for the three different mediator substrates. For 1,4-BQ and DCPIP this is mainly because of a more favourable K_M value. As judged from this kinetic analysis the various ToPOx variants hold promise for an application in biocatalytic processes of for BFC, and here especially ToPOx L545C and T166R seem to be of interest.

The mutations of ToPOx included positions around the catalytic site of POx. T166, Q448, L545, and L547 are part of the substrate-binding domain of POx (residues 50–253, 354–551) [16]. The crystal structure of POx shows a water-filled cavity of about 15,000 Å³. Each monomer of the POx tetramer contains one active site, and the active site entrances face each other as well as the interior solvent-filled void. Any substrate molecule entering the active site first has to pass through one of four channels leading from the surface to the void, and can only then access a narrow active site channel leading to the isoalloxazine ring. A leucine triad (residues 111, 545 and 547) as well as Ser455 and Tyr456 define the entrance of this active-site channel from the void in ToPOx. Replacing one of these leucines in the triad by a cysteine showed the most pronounced effect on the reactivity of ToPOx with the different electron acceptor substrates employed. Currently it is not known how electrons are transferred from the isoalloxazine ring to the electron acceptor substrates employed in our study, but when considering that access to the direct vicinity of the isoalloxazine is controlled by an active-site loop in POx [17,18] and that therefore space is very limited it seems unlikely that these bulky molecules are in direct contact with the isoalloxazine. Thus, a Cys at position 545 at the entrance to the active-site tunnel might participate in the long-range electron transfer from the isoalloxazine to the electron acceptor substrate. Cysteine residues are well known to take part in such electron transfer reactions within a polypeptide [37]. In addition, creating extra space in the active site tunnel might allow the redox mediators such as the rather bulky to get closer to the isoalloxazine, which will improve electron transfer rates as well.

5. Conclusions

The improved kinetics of the POx variant-modified GCEs compared to the WT-GCE suggest the feasibility of fabricating an improved biosensor or BFC, and both L545C and T166R appear to be the most promising ToPOx variants with different mediators for these applications. In general, rapid kinetics experiments with the respective mediator gave good predications about the performance of the individual variants on the electrodes, which is in contrast to steady-state kinetic results obtained for these variants earlier [19]. Although glucose oxidase is by far dominating in the field of bioelectrocatalytic applications, POx shows properties that are beneficial compared to GOx, e.g., a considerably higher catalytic efficiency (k_cat/K_M) for d-glucose, significantly lower Michaelis constants K_M for d-glucose, and reactivity with both anomeric forms of d-glucose [4,38]. Previous bioelectrochemical studies comparing POx and GOx also showed this superiority of POx in certain applications [39]. POx on GCE exhibited a significantly lower K_M in the presence of ferrocene carboxylic acid than GOx-GCE, and the sensitivity of POx-GCE towards glucose on a biosensor was 77.5-fold better than GOx-GCE [40]. The use of 1,4-BQ also improved the sensitivity of another POx-based biosensor [41]. These studies were conducted with wild-type POx, and therefore the beneficial variants described in this study could present attractive alternatives to GOx for bioelectrocatalysis.

Three of the studied mutations introduce a positive charge in the close vicinity of the isoalloxazine. It has been shown previously that positive charges near the FAD can have an effect on the oxidative half-reaction of FAD-dependent oxidoreductases [43]. However, these studies were only focusing on oxygen as electron acceptor, so it is not known at present whether the reaction with alternative electron acceptors will also be affected by the introduction of positive charges. It will certain be worthwhile to look at this in more detail when considering designing engineered variants for bioelectrochemical applications.

Declaration of Competing Interest

The authors confirm that this article content has no conflict of interest.