# Enzymatic biofuel cell on flexible nanoporous gold electrodes

Denise Demurtas, Julia Alvarez-Malmagro, Andrés Felipe Quintero-Jaime, Tanushree Mandal, Sébastien Gounel, Thomas M.B. Reichhart, Anna Lielpetere, Alfons K.G. Felice, Christopher Schulz, Roland Ludwig, et al.

► To cite this version:

Denise Demurtas, Julia Alvarez-Malmagro, Andrés Felipe Quintero-Jaime, Tanushree Mandal, Sébastien Gounel, et al. Enzymatic biofuel cell on flexible nanoporous gold electrodes. Bioelectrochemistry, 2025, 166, pp.109034. 10.1016/j.bioelechem.2025.109034. hal-05136163

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# Enzymatic biofuel cell on flexible nanoporous gold electrodes

Denise Demurtasª, Julia Alvarez-Malmagroª, Andrés Felipe Quintero-Jaimeª,
Tanushree Mandalᵇ, Sébastien Gounelg, Thomas M.B. Reichhartc, d, Anna Lielpetere e,
Alfons K.G. Felice d, Christopher Schulz d, Roland Ludwig c, d, Marcos Pita f, Antonio L. de Lacey f,
Dónal Leech b, Wolfgang Schuhmann e, Nicolas Mano g, Edmond Magner a,*,

## ARTICLE INFO

### Keywords:

Nanoporous gold (NPG)

Enzymatic biofuel cell (EBFC)

Flexible electrodes

Cellobiose dehydrogenase (CDH)

Bilirubin oxidase (BOD)

## ABSTRACT

A glucose/O₂-based biofuel cell employing nanoporous gold electrodes (NPG) supported by Kapton® was prepared. The anode was prepared by drop-casting a solution of Crassicarpon hotsonii cellobiose dehydrogenase (ChCDH) and an Os complex-based polymer and the cathodes by covalent immobilization of Magnaporthe oryzae bilirubin oxidase (MoBOD) on 3-mercaptopropionic acid (MPA) self-assembled monolayer (SAM). Both electrodes were coated with poly(2-methacryloyloxyethyl phosphorylcholine-co-glycidyl methacrylate) (MPC) to reduce biofouling. The anode had a Jmax of 172 ± 10 µA cm⁻² and a KMapp of 19 ± 3 mM in phosphate buffer saline, with a linear detection range from 1 to 5 mM and a sensitivity of 9.4 ± 0.3 µA cm⁻² mM⁻¹. In artificial plasma, the response was saturated at 3 mM, with Jmax of 6.8 ± 10 µA cm⁻², KMapp of 1 mM and a linear detection range from 1 to 5 mM. The cathode had a Jmax of 103 µA cm² and retained 80 % of its response after 18 h of continuous measurement in phosphate buffered saline, while in artificial plasma, the stability was significantly reduced with a half-life of 1 h under continuous operation. The power outputs in PBS and artificial serum were 4.4 and 1.0 µWcm⁻², respectively.

## 1. Introduction

Enzymatic biofuel cells (EBFCs) are electrochemical devices that convert chemical energy into electricity by employing oxidases and reductases as biocatalysts immobilized at the anode and cathode, respectively [1]. The enzymes are selected based on their activity towards biofuels, such as glucose and lactate for the oxidation reaction and oxygen for the reduction reaction; substrates that are widely available in the human body [2]. EBFCs can serve as a sustainable and renewable power source for small devices [3,4] and their power output can be directly correlated to the concentration of a substrate [5]. EBFCs can operate in physiological conditions, such as neutral pH and room temperature, for potential use in a range of applications [1]. However, the design of implantable EBFC involves several challenges regarding the

biocompatibility of the devices, low power density, and limited long-term stability [1]. Various strategies have been investigated to enhance the power output and stability of EBFCs. The electrode material selected is critical and is based on the conductivity, structure and biocompatibility. Nanostructured materials that provide high surface areas, such as multiwall carbon nanotubes (MWCNT) [6], nanoporous gold (NPG) [7] and gold nanoparticles (AuNPs) [8] can enable the use of larger loadings of immobilized enzyme, improving performances in terms of stability and catalytic output [1]. Other strategies for enhancing the performance include enzyme engineering to boost direct electron transfer (DET) and improve substrate selectivity [9], co-immobilization of enzymes and redox mediators for mediated electron transfer (MET) to boost the faradaic output while reducing enzyme's leaching [5]. The inclusion of additional anti-inference layers has been used to mitigate

<footnote>ª Department of Chemical Sciences, Bernal Institute University of Limerick V94 T9PX, Limerick, Ireland</footnote>

<footnote>ᵇ School of Chemistry & Ryan Institute, University of Galway, University Road, Galway, Ireland</footnote>

<footnote>ᶜ Biocatalysis and Biosensing Laboratory, Department of Food Science and Technology, BOKU – University of Natural Resources and Life Sciences, Muthgasse 18, 1190 Vienna, Austria</footnote>

<footnote>d DirectSens Biosensors GmbH, Am Rosenbübel 38, 3400 Klosterneuburg, Austria</footnote>

<footnote>e Analytical Chemistry - Center for Electrochemical Sciences, Faculty of Chemistry and Biochemistry, Ruhr-University Bochum, 44780 Bochum, Germany</footnote>

<footnote>f Instituto de Catálisis y Petroleoquímica, CSIC, c/Marie Curie 2, 28049 Madrid, Spain</footnote>

<footnote>g Centre de Recherche Paul Pascal (CRPP) CNRS, UMR 5031, Univ. Bordeaux, 115 Avenue du Docteur Schweitzer, 33600 Pessac, France</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: Edmond.magner@ul.ie (E. Magner).</footnote>
the influence of interfering species [10].

NPG electrodes are potentially suitable materials in applications where biofouling and stability are critical parameters. Previous reports have described the preparation of NPG on various supports such as polyethyelene terephthalate (PET) [7], glassy carbon electrodes (GCE), glass [11] and polyimide [12]. NPG electrodes can enhance the faradaic response and stability of immobilized redox proteins such as bilirubin oxidase (BOD), glucose oxidase (GOx), cellobiose dehydrogenase (CDH) in comparison to planar electrodes [11,13]. These improvements can be ascribed to the topography of NPGs, which have high surface areas and provide a protected microenvironment for the enzymes. Sensors based on NPG showed enhanced sensitivity, reproducibility, selectivity and long-term stability in human serum samples [14], as well as an enhanced linear detection range for DNA assay [15].

NPGs have also been employed in EBFCs, outperforming planar gold electrodes in terms of power density and stability under pseudo-physiological conditions [1]. Glucose EBFCs use GOx predominantly, with the enzyme catalysing the oxidation of glucose using oxygen as a co-substrate, generating hydrogen peroxide ($H_2O_2$) as a byproduct. However, the accumulation of $H_2O_2$ can inhibit enzyme activity, induce degradation, and amplify inflammation at the tissue level. Some anodes modified with glucose oxidase were coupled with peroxidase [16] or catalase [17] to remove $H_2O_2$. As an alternative, oxygen-independent enzymes, such as GDH or variants of ChCDH, can be employed to avoid this drawback [18]. To date, the most effective EBFC was prepared using GOx from *Aspergillus niger*, and BOD from *Trachyderma tsunodae*, both entrapped in a redox polymer on carbon fibre electrodes modified with CNT. This system had a power output of 740 µW cm⁻² (15 mM glucose in phosphate buffer saline, 37 °C) with an operational stability of 7 h [6]. Previous investigations using ChCDH and MvBOD on NPG reported a power output of 8 µW cm⁻² in 50 mM phosphate-buffered saline containing 5 mM of glucose at 37 °C, at pH 7.4 [19].

Implantable devices must operate effectively in biological fluids containing species, such as ascorbic acid (AA) and uric acid (UA), that can undergo direct oxidation at the electrode surface, acting as an interferent [7]. Moreover, UA has an inhibitory effect on enzymes such as ChCDH [20] and BOD [21]. Non-competitive inhibition of ChCDH by UA and its oxidation products, such as allantoin and allantoic acid, has also been reported [20]. When UA is oxidised in the presence of $O_2$, it binds and extracts $Cu^{2+}$ in the BOD active site, irreversibly deactivating the enzyme [21]. These effects can be mitigated by using anionic polymer coatings such as Nafion® and poly(ester-sulfonic acid) or co-polymers to repel interfering anionic species from the active layer [22]. The above-mentioned approach is often combined with an anti-biofouling coating for measurement in real samples. When sensors are implanted, the foreign body reaction (FBR) occurs leading to encapsulation of the foreign materials by fibrotic tissue [23]. This capsule hinders analytes diffusion rate reducing the performance of the sensor [24]. To mitigate this effect surface coating with zwitterionic polymers such as methacryloyloxyethyl phosphorylcholine (MPC) can reduce the degree of fouling and the thickness of the FBR capsule, improving the overall performance of the sensor [25]. In this study, we describe the preparation of a glucose/$O_2$-based biofuel cell based on the use of nanoporous gold electrodes (NPG) supported by Kapton®. The anode and cathode possessed good stability in PBS, on testing in artificial plasma, both electrodes showed reduced currents and stability, with the stability of the cathode response the limiting factor.

# 2. Experimental

## 2.1. Reagents and solutions

The following reagents were purchased from Sigma-Aldrich: 1,2,2'-ethylenedioxydiethylamine (EDEA, 97.7 %), poly(ethylene glycol) diglycidyl ether Mn 500 and Mn 526 (PEDGE-500 and PEDGE-526), potassium hexacyanoferrate(II) trihydrate, potassium

hexacyanoferrate(III), uric acid (UA, ≥ 99 %), L-ascorbic acid (AA, ≥ 99 %), D-glucose (Glu, ACS reagent), D-fructose (≥ 99 %), D-lactose (≥ 99 %), urea (ACS reagent), l-cystine (≥ 99.7 %) (0.075 mM), $NaH_2PO_4$ (≥ 99 %), $Na_2HPO_4$ (≥ 99 %), calcium sulphate dehydrate (ACS reagent, 98 %), magnesium sulphate (≥ 99.5 %), sodium chloride (≥ 99), sodium bicarbonate (ACS reagent), N-hydroxysuccinimide (NHS) (98 %), 2-(N-morpholino)ethanesulfonic acid hydrate (MES, 99.5 %) and mercapto-propionic acid (MPA) (≥ 99 %), Nafion® 117 solution (~5 % in mixture of lower aliphatic alcohols in water), nitric acid (ACS grade 70 %), hydrogen peroxide in water (30 %, w/w), sulfuric acid (95–98 %), and BSA (≥ 96 %). N-Ethyl-N'-((3-dimethylaminopropyl) carbodiimide hydrochloride (EDC) was obtained from ThermoFisher Scientific. Kapton® 50 HN sheets (50 µm thick) were obtained from Dupont.

Milli-Q® water with a resistivity of 18.2 MΩ cm was supplied by an Elgastat Purelab Pulse system (Elga, UK) and used to prepare all aqueous solutions. Nitrogen (99.99 %) and oxygen gases (99.80 %) were supplied by BOC, Limited. The osmium redox polymer [Os(2,2- bipyridine)₂ (polyvinylimidazole)₁₀ Cl]⁺, denoted as (Os(bpy)₂PVI) was synthesised as described by Mercer et al. [26]. Poly(2-methacryloyloxyethyl phosphorylcholine-co-glycidyl methacrylate (MPC) was synthesised as previously reported [10]. Artificial plasma (AP) was prepared as previously reported in [27].

## 2.2. Enzymes

Cellobiose dehydrogenase from *Crassicarpon hotsonii* (ChCDH) (syn. *Myriococcum thermophilum*) with enhanced activity towards glucose was provided by DirectSens GmbH using a published procedure [28], dissolved in water and stored at -20 °C. The activity was 2.95 ± 0.13 U/mg (300 mM glucose in PBS pH 7.4 at 30 °C, using cytochrome c as electron acceptor). Bilirubin oxidase from *Magnaporthe oryzae* (MoBOD), was isolated, produced and purified as previously reported [29], aliquoted and stored at -80 °C. The enzymatic activity was 546 U/mg (50 mM citrate/phosphate containing 1 mM ABTS at pH of 4 and 37 °C).

## 2.3. Electrode preparation

NPGs were prepared on a Kapton® substrate according to a previous report [12] and dealloyed at 20 °C for 1 and 5 min to immobilize the ChCDH and MoBOD, respectively. Polycrystalline gold electrodes (PG) were cleaned using piranha solution (1 part $H_2O_2$: 3 parts $H_2SO_4$) for 10 min to remove all organic contamination and rinsed with Milli-Q water. PGs were then mechanically polished with 1 µm aluminium oxide and 0.5 µm diamond lapping discs (Mason Technology Ltd.), rinsed with Milli-Q water, and sonicated in a water-ethanol solution for 5 min. Electrochemical cleaning was performed in 0.5 M NaOH solution with a potential window from 0 to -1.5 V vs Ag/AgCl 3 M KCl at 200 mVs⁻¹, for 20 cycles. The electrochemical active surface area (ECSA) was assessed as reported in Section 2.6.

## 2.4. Bioanode

Four different electrode modification protocols were implemented to examine the contribution of individual layers to the performance of the bioanode (Scheme 1). Protocols 1 and 3 (Pr1 and Pr3) did not use MPC and were used as controls, while protocols 2 and 4 (Pr2 and Pr4) included a MPC layer. An additional Nafion® layer was included in Pr3 and Pr4. The active layer on the anode was mixed directly onto the electrode surface using a micropipette, using a mixture of an aqueous suspension of an osmium polymer Os(bpy)₂PVI (5 mg mL⁻¹, 30 µL), ChCDH (0.45 EU, 12.1 µL) and cross-linker PEDGE (15 mg mL⁻¹, 7.4 µL). After allowing the active layer to dry for 3 h in a vacuum chamber at RT, an aliquot of either MPC (10 µL, 5 mg mL⁻¹) (Pr2) or Nafion® (10 µL, 0.5 % in water (wt/v), (Pr3) was applied. The Nafion® layer was applied as described above, followed by deposition of the MPC layer (5 mg mL⁻¹, 18 µL) with EDEA (2.5 mg mL⁻¹, 10 µL) on the Nafion® layer after three
<figure><img src="image_3.png" /><figcaption>**Scheme 1.** Schematic representation of the modified (A) anode and (B) cathode. Created in BioRender. Demurtas, D. (2025) https://BioRender.com/w32h411.</figcaption></figure>

hours (Pr4). Nafion® 0.5 % (wt/v) solutions were freshly prepared at a neutral pH; just before deposition, the solution was purged with N₂ for one minute. All modifications were allowed to dry overnight at 21 °C before measurement.

## 2.5. Biocathode

Before modification, NPG electrodes were characterised as described above, sonicated in H₂O, rinsed and dried under a steam of N₂ and stored under vacuum for one hour to ensure complete dryness. The electrodes were modified by immersion in a 5 mM MPA solution in ethanol overnight at 21 °C and rinsed. An aliquot of MoBOD solution (2.35 mg mL⁻¹, 30 µL) in 10 mM MES buffer was drop-cast onto the electrode surface and left in a vacuum chamber for approximately 10 min to enhance the loading of the enzyme. The electrodes were incubated at 4 °C for one hour. Covalent immobilization of MoBOD was achieved by drop-casting a 40 µL aliquot of a solution of 10 mM MES buffer, pH 6, containing 7.7 mM EDC and 9.5 mM NHS for 90 min at 4 °C on the electrodes [30] (Scheme 1 B). EDC and NHS solutions were prepared just before use to avoid hydrolysis and deterioration of the reagents. After rinsing the electrode with 2 mL of 10 mM MES buffer pH 6, a layer of MPC (18 µL corresponding to 90 µg) and EDEA (10 µL corresponding to 25 µg) or 40 PEDGE (40 µL corresponding to 60 µg) was deposited and allowed to dry overnight at 4 °C.

## 2.6. Electrochemical measurements

Electrochemical studies were performed using a CHI1030 potentiostat (CH Instruments, Austin, Texas). A standard three-electrode cell configuration was used for the characterisation of the electrodes, with a modified NPG as a working electrode (WE), a platinum mesh (ALS Co. Ltd., Tokyo, Japan) as the counter electrode (CE) and a Ag/AgCl (3 M

KCl, IJ Cambria Scientific Ltd., UK) as the reference electrode (RE). In the two-electrode set-up used for the EBFC characterisation, the anode was used as a working electrode (WE), and the cathode was used as a combined CE-RE, separated by a distance of ca. 0.5 cm. The power density was measured in a 10 mL solution of oxygen-saturated phosphate buffer (50 mM phosphate, 5 mM glucose and 150 mM NaCl at pH 7.4, (PBS) and in artificial plasma containing 10 mM of glucose at a potential sweep rate of 1 mV s⁻¹.

The ECSA was determined through electrochemical stripping of gold oxide by applying a potential scan from -0.2 to 1.65 V vs Ag/AgCl at 0.1 Vs⁻¹ in 0.5 M H₂SO₄. The charge associated with the gold oxide peak was used to measure the surface area using a conversion factor of 390µC cm⁻² [31]. The geometric surface area (Ageo) was determined by taking a high-resolution photograph of each NPG electrode on a millimetre grid, followed by analysis with ImageJ software [32].

The properties of the electrodes were examined in PBS and in artificial plasma (AP) prepared as previously described [27]; at 37 ± 1 °C, unless specified otherwise. An aqueous stock solution of glucose (2 M) was prepared, allowed to reach equilibrium for 24 h before all measurements, and stored overnight at RT. A solution of 2.5 mM ferroferricyanide (1:1) in phosphate buffer (PB) 50 mM at pH 7.4 was employed to examine the surface coverage of the MPA self-assembled monolayer (SAM) of the modified NPG electrodes [33]. After conditioning for 30 min in PBS or AP, the response of the bioanode was examined using amperometry (E = 0.4 V vs Ag/AgCl with stirring at 150 rpm) or cyclic voltammetry (CV) over the potential range 0.05–0.4 V vs Ag/AgCl. The biocathode was characterised by purging N₂ (inert gas) or O₂ (substrate) into the electrolyte solution for 20 min to reach saturation, followed by amperometry (E = 0.1 V vs Ag/AgCl with bubbling) or CV 0.7–0.1 V vs Ag/AgCl. Current densities were calculated using the geometric area of the working electrode of ca 0.25 cm². The coverage of Os polymer (Γ) was estimated from the peak charges obtained using CV
at a slow scan rate as reported previously [12].

The kinetic parameters, $J_{max}$ and $K_{app}^{app}$, were determined using a nonlinear regression Michaelis and Menten model (Eq. 1) and the limit of detection was determined using Eq. 2 [12].

$$J = \frac{J_{max}[S]}{K_{app} + [S]} \quad (1)$$

$$LOD = \frac{3.3 \times \sigma_{blank}}{\text{slope}} \quad (2)$$

The error bars represent the standard deviation of n = 3 electrodes.

# 3. Results and discussion

## 3.1. Morphological analysis

NPG electrodes were prepared on a Kapton® support using the dealloying conditions described. Electrodes dealloyed in HNO₃ for 1 and 5 min at 20 °C had average pore diameters of 17 and 15 nm, with surface roughness ($R_f$) of 16 ± 4 and 6 ± 1, respectively [12]. These pore sizes, measured from SEM images using ImageJ [34] are compatible with the dimensions of the ChCDH (dimensions of FAD domain~7.2 × 5.7 × 4.5 nm and cyt-c domain 3.0 × 3.6 × 4.7 nm) [9] and of MoBOD based on the homologous structure of *Myrothecium verruccaria* BOD (4 × 5 × 6 nm) [35].

## 3.2. ChCDH modified bioanode

ChCDH is a dehydrogenase containing a FAD cofactor and a cyt-c domain with a heme b connected through a flexible linker. ChCDH can

function via direct (DET) and mediated electron transfer (MET) through the cyt-c and FAD domains, respectively [36]. This study used Os (bpy)₂PVI as a redox mediator for ChCDH on NPG to enable MET from the FAD domain.

Cyclic voltammograms of Os(bpy)₂PVI (Fig. 1A) in the absence of glucose displayed a quasi-reversible response typical of the $Os^{3+}/Os^{2+}$ redox couple with a value of $E_{1/2}$ of 250 mV, ca. 30 mV more positive than the literature value [37]. A surface coverage of 3.35 ± 0.94 nmol cm⁻² was recorded, indicative of the presence of multiple layers on the electrode [37]. The redox process can be described by a surface-confined process at slow scan rates (5–20mV s⁻¹) (Fig. 1 B), switching to a diffusion-controlled process at higher scan rates (Fig. 1 C) [38]. On addition of glucose ($v = 1$ mV s⁻¹), a sigmoidal response characteristic of a catalytic response was observed (Fig. 1 D), demonstrating effective MET between ChCDH and the redox active sites, in agreement with previous findings [36].

The properties of redox hydrogels depend on the polymer used and the degree of crosslinking achieved; high levels of crosslinking can reduce the hydration of the polymer unless the tether linking the redox centres is sufficiently long to enable electron diffusion/hopping [39]. Two different cross-linkers, PEDGE (500 and 526 units), were used to assess which would provide more efficient electron transfer while minimising leakage of the redox moieties [40]. A higher $J_{max}$ of 224 ± 5 µA cm⁻² was achieved using PEDGE.

500 as cross-linker (Fig. 2 A), which has a more dense crosslinking of the redox polymer, as reported by Heller [41], than PEDGE-526 ($J_{max}$ of 183 ± 4 µA cm⁻²). There was no significant variation in $K_{app}^{app}$ with values of 16 ± 1 and 14 ± 0.9 mM for PEDGE-500 and PEDGE-526, respectively. A linear response range of 0–5 mM was obtained with both PEDGEs with sensitivities of 12.6 ± 1.0 (PEDGE-500) and 10.1 ± 0.8 µA

<figure><img src="image_4.png" /><figcaption>**Fig. 1.** (A) Cyclic voltammograms of NPG/ChCDH/Os(bpy)₂PVI/PEDGE-500/MPC as a function of scan rate ($v$); Plots of peak current densities vs. (B) $v$ and (C) $v^{1/2}$. Conditions: PBS at room temperature. (D) Cyclic voltammograms ($v = 1$ mV s⁻¹) of ChCDH/Os(bpy)₂PVI/PEDGE-500/MPC at increasing concentrations of glucose. Conditions: PBS at 37 °C.</figcaption></figure>
<figure><img src="image_5.png" /><figcaption>**Fig. 2.** (A) Baseline corrected amperometric response as a function of glucose concentration for electrodes (n = 3) modified with ChCDH (0.45 U), Os(bpy)₂PVI (150 µg), PEDGE-500 or PEGDE-526 (111 µg) and MPC (100 µg) and (B) corresponding plot of current versus concentration of glucose. Measurement conditions: E = 0.4 V, PBS at 37 °C with stirring at 150 rpm.</figcaption></figure>

cm⁻² mM⁻¹ (PEDGE-526) and LOD of 1.97 and 0.58 mM, respectively.

Higher values of Jmax and Kapp and higher coverage of Os polymer (Table 1) were obtained with PEDGE-500. When this electrode was modified with the zwitterionic polymer MPC the values Jmax and Kapp were 153 ± 4 µA cm⁻² and 14 ± 0.9 mM, respectively with a sensitivity of 8.5 ± 0.6 µA cm⁻² mM⁻¹ and a LOD of 2.37 mM. Electrodes without MPC had a higher Jmax of 224 ± 5 µA cm⁻² and a higher sensitivity of 12.6 ± 1 µA cm⁻² mM⁻¹ despite the fact that the MPC modified electrodes had almost double the coverage of Os species, 3.4 ± 0.9 vs 6 ± 3 mmol cm⁻². The observed decrease in catalytic current can be ascribed to increased diffusional limitations for glucose in the matrix [42]. The sensitivity of the MPC modified electrode decreased by 19%, in contrast to a 1.5-fold increase previously described on graphite. In the latter case, the increase was ascribed to an increased rate of diffusion of glucose through the polymeric layer. The values of Jmax of the electrode with and without MPC were ca. 40% of that on graphite and the values of Kapp were half of the reported value [43]. However, a direct comparison is not possible as the enzyme loadings were not determined. The density and thickness of the polymer/enzyme/ layer on NPG plays an important role in determining the catalytic response. On electrodes modified with same osmium polymer complex and lactate dehydrogenase, thinner layers led to an increased response, while thicker layers resulted in clogged pores, limiting mass transport through the film [7,44]. In this study, the addition of an extra MPC layer, which on visual inspection is full mixed with the osmium polymer, may result in slower rates of diffusion of glucose arising from the increased thickness of the polymer layer.

In physiological fluids, the effect of interferents such as AA and UA, which can be directly oxidised on the electrode surface and the presence of potential non-competitive inhibitors such as UA, chlorides and urea [8] must be considered. UA can act as an uncompetitive inhibitor of ChCDH, reducing the enzymatic activity by up to 50% [20,45]. An

anionic perm-selective membrane, such as Nafion®, that is permeable to glucose but repels anionic species such as AA and UA [42], can mitigate this effect. The response of the electrodes (Fig. 3A and Table 2) in the presence and absence of Nafion® showed a Jmax of 171 ± 10 µA cm⁻² and Kapp of 13 ± 2 mM, indicating that the additional polymer layer did not significantly restrict the rate of diffusion of glucose.

The surface coverage of the Os polymer on addition of MPC and Nafion® (Table 2) was higher with the optimal coverage obtained by first coating the enzyme layer with Nafion® followed by MPC, to yield a final coverage of 6.7 nmol cm⁻². This improvement can be attributed to the electrostatic interaction between the positively charged Os(bpy)₂PVI and the negatively charged sulfonate groups in Nafion®, which helps mitigate leakage of the redox moieties by forming a physical barrier over the electrode [46]. This electrode also displayed the highest value of Jmax (244 µA cm⁻²), however with a decrease in sensitivity, the origin of which is unclear.

The linear range increased to 9 mM for Os(bpy)₂PVI/ChCDH/ PEDGE-500/Nafion®/MPC and can be ascribed to the increased diffusional effects of the additional layers (Table 2). Kapp was not significantly affected by presence of additional layers but was approximately half of the value reported in the literature for the same polymer layers on graphite [36].

The electrocatalytic activity of the bioanode was continuously monitored for 18 h in PBS and AP (Fig. 4), with a stable response obtained for both coated and uncoated electrodes for up to 6 h. The uncoated electrodes showed a decrease in activity of ca. 50% after 18 h. In PBS, NPG/Os(bpy)₂ PVI/ChCDH/PEDGE-500 and NPG/Os(bpy)₂ PVI/ ChCDH/PEDGE-500/MPC retained 27.5% (half-life >12 h) and 69% current output after 18 h. CVs obtained before and after 18 h of continuous glucose oxidation in PBS showed that a catalytic response was still present for uncoated and MPC-coated electrodes (Fig. S1). However, in AP, the half-lives of both electrodes decreased significantly to 3 h, with no detectable response after 6 h. The response of the polymer layer was tested before and after amperometric measurement in both PBS and AP; the characteristic Os redox peaks, prior to glucose addition, were visible at 250 mV vs Ag/AgCl (Fig. S1 A and B). After 18 h of continuous measurement in AP, a significant loss in the response of the redox polymer was observed, equally for the uncoated and the MPC coated electrodes, probably as a result of the leaching of the Os redox complexes [47,48] with the result that a calibration curve could not be obtained (Fig. S1 B).

Amperometric calibration in AP showed poor electrochemical performance at a Nafion®-MPC coated electrodes (Fig. S2), reaching saturation at 5 mM and a Jmax value of 6 µA cm⁻² while no response was

<table><caption>**Table 1**

List of kinetics parameters for bioanode using different cross-linkers in PBS.</caption><thead><tr><th>Protocol</th><th>K<sup>app</sup><sub>M</sub><br/>Mm</th><th>J<sub>max</sub>/<br/>µA<br/>cm<sup>-2</sup></th><th>Sensitivity/<br/>µA cm<sup>-2</sup><br/>mM<sup>-1</sup></th><th>LOD<br/>mM</th><th>Γ/<br/>nmol<br/>cm<sup>-2</sup></th><th>Linear range<br/>mM</th></tr></thead><tbody><tr><td>PEDGE-<br/>500</td><td>16 ±<br/>1</td><td>224<br/>± 6</td><td>12.6 ± 0.9</td><td>1.97</td><td>3.4 ±<br/>0.9</td><td>0-5</td></tr><tr><td>PEDGE-<br/>526</td><td>14 ±<br/>0.9</td><td>183<br/>± 4</td><td>11 ± 0.8</td><td>0.58</td><td>2.8 ±<br/>0.5</td><td>0-5</td></tr><tr><td>PEDGE-<br/>500 +<br/>MPC</td><td>14 ±<br/>0.9</td><td>153<br/>± 4</td><td>9.4 ± 0.6</td><td>2.37</td><td>6 ± 3</td><td>0-5</td></tr></tbody></table>
<figure><img src="image_6.png" /><figcaption>**Fig. 3.** (A) Baseline corrected amperometric response as a function of glucose concentration for electrodes (n = 3) modified with ChCDH (0.45 U), Os(bpy)₂PVI (150 µg) and PEDGE (111 µg) and subsequently with MPC and Nafion®; (B) plots of current density versus the concentration of glucose. Measurement conditions: MPC (100 µg) or Nafion® (10 µL of 0.5 % (wt/v), or Nafion® (10 µL of 0.5 % (wt/v)) with MPC (90 µg) EDEA (25 µg). E = 0.4 V, PBS at 37 °C with stirring at 150 rpm.</figcaption></figure>

<table><caption>**Table 2**

List of kinetic parameters of the bioanode in PBS.</caption><thead><tr><th>Protocol</th><th>K<sub>M</sub><sup>app</sup><br/>/<br/>mM</th><th>J<sub>max</sub><br/>/µA<br/>cm<sup>-2</sup></th><th>Sensitivity<br/>/µA cm<sup>-2</sup><br/>mM<sup>-1</sup></th><th>LOD/<br/>mM</th><th>Γ /<br/>nmol<br/>cm<sup>-2</sup></th><th>Linear<br/>range /<br/>mM</th></tr></thead><tbody><tr><td>Os(bpy)<sub>2</sub>PVI/<br/>ChCDH/<br/>PEDGE-500</td><td>16<br/>± 0.9</td><td>222<br/>± 6</td><td>12.7 ± 0.9</td><td>1.76</td><td>3.4 ±<br/>0.9</td><td>0-4</td></tr><tr><td>Os(bpy)<sub>2</sub>PVI/<br/>ChCDH/<br/>PEDGE-500/<br/>MPC</td><td>15<br/>± 1</td><td>155<br/>± 4</td><td>9.3 ± 0.7</td><td>2.31</td><td>6 ± 3</td><td>0-4</td></tr><tr><td>Os(bpy)<sub>2</sub>PVI/<br/>ChCDH/<br/>PEDGE-500/<br/>Nafion®</td><td>13<br/>± 2</td><td>171<br/>± 10</td><td>12.0 ± 1.2</td><td>0.07</td><td>5.6 ±<br/>0.1</td><td>0-3</td></tr><tr><td>Os(bpy)<sub>2</sub>PVI/<br/>ChCDH/<br/>PEDGE-500/<br/>Nafion®<br/>/MPC</td><td>17<br/>± 1</td><td>244<br/>± 7</td><td>8 ± 0.5</td><td>1.24</td><td>6.7 ±<br/>0.5</td><td>0-9</td></tr></tbody></table>

evident on MPC modified electrodes. The reduction of the catalytic activity of ChCDH can be attributed to uncompetitive inhibition by UA [20], AA and urea [49]. While electrodes without a Nafion® coating still

showed catalytic activity after conditioning in AP (data not shown), no detectable amperometric response was observed, probably indicating that Nafion® is not as effective in preventing biofouling as MPC [43].

### 3.3. BOD modified biocathode

Biocathodes were prepared using PG and NPG@Kapton® electrodes modified with a self-assembled monolayer of MPA for covalent immobilization of the multicopper oxidase MoBOD. The enzyme shows high catalytic activity at neutral pH with low levels of inhibition by urea [29]. MPA is a short-chain thiol that promotes DET for bilirubin oxidases, enabling a more favourable orientation of the MoBOD T1 site with its -COO⁻ terminal groups. The T1 site enables direct electron transfer with the electrode, while the T2/T3 sites catalyse the reduction of O₂ to H₂O [50]. Moreover, the spacer effect of MPA prevents direct contact between the enzyme and the Au surface, avoiding rapid denaturation [51]. The integrity of the SAM was assessed by examining phosphate adsorption and desorption peaks at -0.1 and -0.2 V vs Ag/AgCl 3 M KCl [33] (Fig. S3) and by using ferro: ferricyanide to probe the level of surface coverage (Fig. S4). The electrochemical response of ferricyanide at the bare electrodes was quasi-reversible, with a peak separation close to 59 mV. On modification of the electrodes with thiols, lower current densities and larger peak separations (> 59 mV) were observed at both

<figure><img src="image_7.png" /><figcaption>**Fig. 4.** Operational stability of electrodes (n = 3) modified with ChCDH (0.45 U), Os(bpy)₂PVI (150 µg), PEDGE (111 µg) with (red lines) and without (black lines) an MPC (100 µg) coating in (A0 PBS and (B) artificial plasma. Measurement conditions: E = 0.4 V, 5 mM glucose at 37 °C under stirring at 150 rpm. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>
electrodes (Fig. S4 A and B), indicating that the surface was not completely covered, with coverages of 60.6 and 178.9 nmol cm⁻², for PG and NPG electrodes, respectively, assessed by stripping voltammetry at basic pH (Fig. S5). The typical structures of SAMs of short chain alkane thiols possess defects that partially allow the redox probes to reach the underlying surface; this effect is more pronounced in the presence of edges and defects on NPG [52].

The response of MoBOD on bare PG using PEDGE-500 as a cross-linker showed an electrocatalytic current of 5 µA cm⁻² (Fig. S6 A). When the Au surface was modified with an MPA monolayer, the current increased over 12-fold; this behaviour is attributed to the more favourable orientation of the T1 site towards the electrode surface, which facilitates DET. Electrodes modified with MPA were also tested using EDC/NHS coupling of the primary amines on the protein surface to the -COO⁻ groups of the SAM (Fig. S6 B) to obtain a monolayer of the enzyme. The protocols employing the two cross-linkers showed similar current densities of ca. 60 µA cm⁻², indicating that the amount of electroactive immobilized MoBOD was not dependent on the nature of the cross-linker.

The catalytic response of the enzyme using PEDGE-500 and the EDC-NHS as cross-linkers on NPG gold electrodes dealloyed at 20 °C for 1 min was examined using cyclic voltammetry in PBS (Fig. 5 A). When tested in AP, a coating of MPC crosslinked with EDEA was drop-cast on the electrodes to reduce biofouling. A maximum catalytic current of 259.1 µA cm² was obtained in PBS (Fig. 5 A), a four-fold increase when compared to the response on PG. The hysteresis observed (Fig. 5) at approximately 250 and 400 mV after three potential cycles can be attributed to incomplete activation of the enzyme [49], visible in the third potential scan after 1 and 2 h at more positive potentials. Hysteresis has been reported on nanostructured gold [51] and spectrographic graphite [50] electrodes and attributed to the switching of the enzyme from a resting oxidised (RO) form to an alternative resting form (AR) in the presence of chloride (150 mM) to a fully reduced and active form able to reduce oxygen to water at a potential of 0.1 V. Lower potentials were not investigated to avoid oxygen reduction which can cause detachment of the SAM [53]. When using cyclic voltammetry, the stability of the response was relatively poor; after one hour (three CVs per hour with storage in PBS at 37 °C at open circuit potential), only 20 % of activity remained. The response obtained on addition of NaF (15 mM), which would completely inhibit the activity of BOD, overlapped with the voltammogram obtained after 17 h Fig. 5 A).

Electrodes modified with EDC-NHS and PEDGE-500 as cross-linkers (Fig. 5 B) tested through chronoamperometry had current densities of 103 and 106 µA cm² (E = 0.15 V). In both cases, ca. 80 % of electrocatalytic activity was retained after 18 h of continuous measurement,

demonstrating the stability of the immobilized MoBOD in 50 mM PBS, pH 7.4 at 37 °C in the presence of 150 mM of chloride. When the activity of the enzyme was tested in solution (100 mM phosphate-citrate buffer, pH 7 at 37 °C in the absence of inhibitors) [29] a half-life of 5 h was obtained, indicating that the enzyme-modified electrode was substantially more stable. MoBOD has been shown to undergo DET on macro-structured gold [54] and on carbon substrates such as GCE modified with carbon nanotubes and MWCNT [55] with stability that ranged from 20 h to several days. However, direct comparisons with the electrodes used here are not feasible as the conditions vary significantly (composition of buffer and salts, pH, temperature, measurement techniques and storage conditions). For instance, MoBOD adsorbed on GC/MWCNT, retained up to 73 % of its activity for 24 h at room temperature in 100 mM McIlvain buffer, pH 6 in the absence of chloride. When the response of MoBOD adsorbed on GC/MWCNTs was examined in O₂-saturated 200 mM phosphate 100 mM citrate buffer, 50 % of the activity was retained after two days (response measured daily storage in deoxygenated wet conditions at 4 °C) [55]. The response of a macroporous gold-based EBFC using Cys-MoBOD and GOx was monitored in PBS (137 mM NaCl) and in subcutaneous tissue and was stable for 20 h under both conditions [54]. MoBOD on diazonium modified GCE and CN-MWCNT retained 75 % activity for 5 days in buffer (100 mM phosphate buffer, pH 6) [56]. The comparison of our system to these results shows that the designed biocathode can operate in pseudo-physiological conditions, even under non-ideal conditions.

The response of the NPG/MPA/MoBOD/EDC-NHS/MPC electrode in AP showed a catalytic current of 50 µA cm⁻², with a half-life of 1 h and a complete loss in response after 6 h (Fig. S7). The crosslinked MPC should mitigate detachment of loosely bound enzyme species and as inactivation by chloride is more evident at acidic pH [57], the decrease in response is more likely affected by the presence of UA [21] and AA [51]. The immobilized enzyme retained 25 % activity after 2 h with a current density of 15 µA cm⁻², a response sufficient to meet the current requirement of the anode, which had a longer half-life of 3 h in AP but a lower catalytic current of 10 µA cm⁻². The higher viscosity of AP [32] combined with the lower activity of MoBOD in the media, limits the current output of the cathode as well as its stability.

### 3.4. Glucose biofuel cell

The glucose biofuel cell was tested using NPG/ChCDH/Os(bpy)₂PVI/PEDGE-500/MPC (EBFC1) and NPG(ChCDH/Os(bpy)₂PVI/PEDGE-500/Nafion®/MPC/EDEA (EBFC2) as the anode and NPG/MPA/MoBOD/EDC-NHS/MPC as the cathode. In 5 mM glucose (PBS), EBFC1 possessed an open circuit voltage (OCV) of 495 mV. The polarization curve (Fig. 6

<figure><img src="image_8.png" /><figcaption>**Fig. 5.** Cyclic voltammograms of NPG/MPA/MoBOD/EDC-NHS as a function of time (A). The cell was kept at OCP when the experiment was paused. (B) Amperometric response of NPGs/MPA/MoBOD/EDC-NHS/MPC and NPG/MPA/MoBOD/PEDGE/MPC. Conditions: PBS at 37 °C, saturated with O₂, scan rate 10 mV s⁻¹. Baseline obtained using (A) NaF and (B) N₂ saturation.</figcaption></figure>
A) had a maximum power density of 4.4 µWcm⁻² at 0.180 V and a maximum current density of 37.7 µA cm⁻². A 50% reduction in response after 1 h was observed, in agreement with the stability data obtained for the cathode. When the anode was coated with both Nafion® and MPC (EBFC2), the cell had a lower OCV of 240 mV in PBS with a maximum power density of 1.4 µW cm⁻² (E = 0.11 V) and a maximum current density of 32.2 µA cm⁻². The differences observed between EBFC1 and EBFC2 in PBS (Fig. 6 A) can be attributed to mass diffusion limitations introduced by the additional Nafion® layer in EBFC2. These limitations were not evident during the calibration of the anode in the three-electrode configuration (E = 0.4 V vs Ag/AgCl), where the solution was stirred, likely enhancing the rate of diffusion of glucose to the electrode surface. This hypothesis is confirmed by the decrease in the OCV value, which was half of that obtained with EBFC1 and is indicative of an increase in the resistance.

In AP, EBFC1 and EBFC2 had OCV of 367 mV and 200 mV, respectively, with maximum current densities of 14.9 and 23.4 µA cm⁻² and maximum power densities of 1.02 and 0.35 µW cm⁻² at potentials of 137 and 58 mV, respectively. The two biofuel cells possessed about 40 % of the current density obtained in PBS, while the power outputs of EBFC1 and EBFC2 decreased by 25 % and 50 %, respectively.

The reduction of ca. 100 mV in OCV for both cells in AP, can be explained by the presence of inhibitors, which affects the onset potential of MoBOD and reduces the catalytic current [7]. Stability control experiments of anodes uncoated or coated with MPC tested for 18 h in AP showed a significant loss of Os complex and minimal enzymatic activity (Fig. 4 and Fig. S1). The loss of Os polymer and the reduced performance of the biocathode in AP explain the rapid loss of power of the EBFC in this medium. Cadet et al. reported a 45 % decrease in power density loss in buffer and 65 % in blood using an MET-based EBFC employing GDH and MoBOD, with the losses primarily ascribed to leakage of Os species with only 20 % loss from the presence of inhibitors [47]. Electrografting amines [24] or aryl diazonium salts [56] to carbon or gold electrodes could significantly improve the adhesion of redox polymers and will be the object of future work.

Previous studies employing ChCDH and BOD in human serum [57], blood [8] and in vivo in rats [58] showed power outputs of 4, 3 and 2 µW cm⁻² with OCVs of 0.58, 0.63 and 0.40 V, respectively, and half-lifes of 2 h using a range of electrode configurations. The OCV obtained for EBFC1 (367 mV) in AP is comparable to previous results. Moreover, the anode and cathode were coated with crosslinked MPC; this additional layer should help decrease the level of detachment of the enzyme while reducing biofouling. The Nafion® coating on the bioanodes should reduce the effect of interferents, as shown in testing the coated electrodes in AP. However, EBFC2 showed a lower power output and stability than electrodes coated only with MPC (Fig. 7), probably because

the reduced potential difference between the anode and cathode is indicative of an increased resistance in the system, as was also observed in PBS. After 30 min of stabilization, the power output of EBFC1 decreased from 4 to 1 µW cm⁻² (Fig. 7) followed by a stable power output for a period of 3 h, retaining 20 % of the original power over this time period. In contrast, EBFC2 exhibited a 20-fold lower power output (0.20 µW cm⁻²) at the beginning of the measurement, which declined by 50 % after ca. 30 min with a complete loss of power after 1 h (Fig. 7). These results indicate that EBFC1 has a higher power output than EBFC2; nevertheless, further investigation is required to improve the performance of the anode and the stability of the cathode, especially in complex media.

## 4. Conclusions

In this work, the development of a membrane-less biofuel cell based on a flexible NPG electrode supported by Kapton® has been described for potential application as an implantable device. The bioelectrodes were modified with an O₂-insensitive ChCDH and an Os polymer to

<figure><img src="image_9.png" /><figcaption>Fig. 7. Power output of EBFC1 and EBCFC2 in AP at 37 °C, 5 mM glucose at applied potentials +0.137 and 0.05 V, respectively (n = 3 electrodes).</figcaption></figure>

<figure><img src="image_10.png" /><figcaption>Fig. 6. Power density (red) and polarization curves (black) as a function of cell voltage obtained for BFCs consisting of a NPG/MPA/EDC-NHS/MoBOx cathode combined with (A) NPG/ChCDH/Os(bpy)₂PVI/MPC and (B) NPG ChCDH/Os(bpy)₂PVI/Nafion®/MPC anodes. Curves obtained in PBS (full lines) and AP (dotted lines) containing 5 mM of glucose at 37 °C. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>
achieve MET and MoBOD covalently attached to a SAM for DET; both designs were coated with a zwitterionic polymer (MPC). Phosphate buffer saline and artificial plasma were used as the electrolyte solutions. The bioanode, ChCDH/Os(bpy)₂PVI/PEDGE/MPC displayed Michaelis and Menten behaviour with $J_{max}$ of $155 \pm 4$ µA cm⁻², $K_{M}^{app}$ of $14.5 \pm 1$ mM and linear detection ranges of 1–4 mM, and the biocathode MPA/MoBOD/MPC a $J_{max}$ of $103$ µA cm⁻² with a stability up to 18 h respectively. The assembled enzymatic biofuel cell EBFC1 generated a power of $4.4$ µWcm⁻² at 180 mV versus RE in buffer. In artificial plasma, the performance was reduced to a $J_{max}$ of $6.8 \pm 10$ µA cm⁻² and a $K_{M}^{app}$ of 1 mM with a linear detection range from 1 to 3 mM; the biocathode instead showed a $J_{max}$ of $50$ µA cm⁻², while the corresponding EBFC generated a power output of $1$ µWcm⁻² at a potential of 137 mV, for up to 3 h in artificial plasma. When Nafion® was added to the bioanode prior to MPC to reduce the impact of anionic inhibitors in PBS, $J_{max}$ improved to $244 \pm 4$ µA cm⁻², and $K_{M}^{app}$ of $17 \pm 1$ mM was similar to the previous response. However, EBFC2 performed less well than EBFC1 both in PBS and AP. The response was limited by the cathode's operational stability in AP, possibly hindered by inhibition by UA, AA oxidation and an overall increased system resistance. The biofuel cell is based on a flexible NPG support which lends itself for use in implantable systems, however additional investigation of alternative perm-selective polymers and grafting strategies are required to improve the stability.

### CRediT authorship contribution statement

**Denise Demurtas:** Writing – original draft, Visualization, Validation, Methodology, Investigation. **Julia Alvarez-Malmagro:** Investigation. **Andrés Felipe Quintero-Jaime:** Investigation, Conceptualization. **Tanushree Mandal:** Investigation, Conceptualization. **Sébastien Gounel:** Investigation. **Thomas M.B. Reichhart:** Investigation. **Anna Lielpetere:** Investigation. **Alfons K.G. Felice:** Resources, Investigation. **Christopher Schulz:** Resources, Investigation, Funding acquisition. **Roland Ludwig:** Investigation, Funding acquisition, Conceptualization. **Marcos Pita:** Investigation, Funding acquisition, Conceptualization. **Antonio L. de Lacey:** Investigation, Funding acquisition, Conceptualization. **Dónal Leech:** Resources, Investigation, Funding acquisition, Formal analysis, Conceptualization. **Wolfgang Schuhmann:** Resources, Investigation, Funding acquisition, Conceptualization. **Nicolas Mano:** Resources, Methodology, Conceptualization. **Edmond Magner:** Writing – review & editing, Supervision, Resources, Project administration, Investigation, Funding acquisition, Formal analysis.

### Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

### Acknowledgements

This work has received funding from the European Union's Horizon 2020 research and innovation MSCA ITN program under grant agreement N°813006, IMPLANTSENS and the MSCA IF program under grant agreement N°101024374, PATTENZYME.

### Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.bioelechem.2025.109034.

### Data availability

Data will be made available on request.

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