# Expanded "Popcorn" Graphite bioelectrocatalytic electrode materials for glucose/O2 biofuel cells: A cost-efficient alternative to nanostructured carbons

Paulo Henrique M. Buzzetti, Anurag Mohanty, Pierre Armand, Andrew J.

Gross, Yannig Nedellec, Michael Holzinger

► To cite this version:

Paulo Henrique M. Buzzetti, Anurag Mohanty, Pierre Armand, Andrew J. Gross, Yannig Nedellec, et al.. Expanded "Popcorn" Graphite bioelectrocatalytic electrode materials for glucose/O2 biofuel cells: A cost-efficient alternative to nanostructured carbons. Carbon, 2025, 244, pp.120691. 10.1016/j.carbon.2025.120691. hal-05261774

HAL Id: hal-05261774

https://hal.science/hal-05261774v1

Submitted on 15 Sep 2025

HAL is a multi-disciplinary open access
archive for the deposit and dissemination of sci-
entific research documents, whether they are pub-
lished or not. The documents may come from
teaching and research institutions in France or
abroad, or from public or private research centers.

L'archive ouverte pluridisciplinaire HAL, est
destinée au dépôt et à la diffusion de documents
scientifiques de niveau recherche, publiés ou non,
émanant des établissements d'enseignement et de
recherche français ou étrangers, des laboratoires
publics ou privés.

<figure><img src="image_1.png" /></figure>

Distributed under a Creative Commons Attribution 4.0 International License
# Expanded “Popcorn” Graphite bioelectrocatalytic electrode materials for glucose/O₂ biofuel cells: A cost-efficient alternative to nanostructured carbons

Paulo Henrique M. Buzzetti<sup>a,b</sup>, Anurag Mohanty<sup>a,b</sup>, Pierre Armand<sup>a</sup>, Andrew J. Gross

<sup>a,b</sup>, Yannig Nedellec<sup>a,b</sup>, Michael Holzinger<sup>a,b*</sup>

<sup>a</sup> Univ. Grenoble Alpes, DCM UMR 5250, F-38000 Grenoble, France.

<sup>b</sup> CNRS, DCM UMR 5250, F-38000 Grenoble, France

## Abstract

Here we demonstrate the beneficial use of expanded graphite as an electrode material suitable for enzyme wiring for bioelectrocatalytic energy conversion. Expanded graphite is obtained via microwave treatment using commercial expandable graphite, which leads to a drastic popcorn-like volume increase. After an adjusted dispersion and filtration procedure, enzymatic bioanodes are formed using FAD-glucose dehydrogenase (FAD-GDH) with 9,10 phenanthroline-5,6-dione (PLQ) as an electron shuttle. At the biocathode, heme is used as a promoter to immobilize and orientate bilirubin oxidase (BOD). The expanded popcorn graphite (EPG) electrodes are characterized with electron microscopy, contact angle, and Raman spectroscopy. Its surface area, determined with the Brunauer-Emmett-Teller (BET) method, is about ten times lower than lab-made carbon nanotube buckypapers (CNT-BPs). However, the electroactive surface area revealed comparable values for both materials, within the same order of magnitude. The bioelectrocatalytic performances were determined with cyclic voltammetry. For the bioanode, a current density of 2.87 mA cm⁻² is obtained at 100 mmolL⁻¹ glucose concentration at 0.3 V vs Ag/AgCl (sat. KCl). The biocathode provided a current density of ~0.41 mA cm⁻² in oxygen saturated buffer solution at 0.2 V. These performances are compared with lab-made CNT-BPs and commercial CNT-BPs. Finally, a complete glucose biofuel cell was formed that produced a maximum power density of 0.15 mW cm⁻² at an open circuit voltage of 0.41 V.

## Keywords:

Expandable graphite; expanded “popcorn” graphite; carbon-based electrode; bioelectrocatalysis; enzymatic biofuel cells

<footnote>* Corresponding author. E-mail address: *michael.holzinger@univ-grenoble-alpes.fr</footnote>
# 1. Introduction

Enzymatic biofuel cells are widely considered as a purely academic research application due to the low power production and limited operational life time [1]. However, related to the vast evolution of low power consuming electronics for single or short period use, enzymatic biofuel cells gained high interest in the industrial sector as eco-friendly alternatives to currently used button batteries to power these devices [2-4]. Biofuel cells operate in a similar way to classic fuel cells, relying on chemical to electrical conversion. Unlike fuel cells, the catalysts are biomacromolecules (enzymes) instead of noble metals or metal alloys. The use of enzymes provides attractive advantages relating to their optimal activity at room temperature and in neutral aqueous solutions, their turnover, and their high specificity which permit energy harvesting in complex media without the need for ion-conducting membranes [5]. The challenge with employing enzymes is to establish efficient electron transfer(s) between the active site and an electrode. Many strategies are available to achieve this and are distinguished by two modes: direct electron transfer (DET) and, when necessary, mediated electron transfer (MET) [6, 7]. Mediated electron transfer relies on an electron shuttle, generally a small redox active molecule, to enables such electron transfers. To reach the necessary energy demands for practical applications, nanomaterials have become essential components. For example, nanomaterials can optimize the electrocatalytic performances in terms of electron transfers at the highest possible cell voltages. Among such nanomaterials, carbon nanotubes (CNTs) became the material of choice due to the possibility to formulate electrode interfaces with beneficial high surface area and attractive pore structures and inertness. One possibility is to form freestanding macroscopic electrodes of different shapes ranging from fibres to papers (e.g. so-called carbon nanotube buckypapers (CNT-BPs)) to pellets [7, 8]. The development of enzymatic biofuel cells has remained at an academic level due to limitations in terms of stability and activity, but also due to the relatively high costs of CNTs and concerns about their suspected
toxicity [9-11]. Additionally, recent legal requirements addressing potential health hazards associated with nanosized materials is a constant drawback for industrial developments [12, 13].

Expandable graphite was first reported in the 19th century and was referred to as “graphite salts” [14]. This expandable graphite is generally obtained by thermal treatment of mineral graphite with strong inorganic acids and oxidizers [15, 16]. Sulfuric acid has a great capacity to intercalate in between the graphene sheets during this treatment, leading to graphite sulphate or expandable graphite. The expansion is initiated via thermal or microwave treatment that provokes spontaneous evaporation of the intercalants, causing partial exfoliation of the different graphene layers [17], which then leads immediately to a tremendous volume increase similar to the formation of popcorn from kernels.

Such expanded popcorn graphite (EPG) has been used for decades in composite materials such as flame-retardants, lubricants, additives for heat and energy storage, and is even found in pencil graphite [18-21]. More recently, EPG was reported to be an excellent precursor for the large-scale synthesis of high-quality monolayer graphene [22]. However, there are to date surprisingly few examples concerning the use of EPG as an electrode material for bioelectrochemical applications, despite its vast use in industrial products. There are just two reports where EPG was used to construct anodic electrodes for microbial fuel cells, but the authors did not discuss the benefits or report the bioelectrochemical performances of the EPG electrodes [23, 24]. This minimal research activity can be explained by the fact that freshly prepared EPG is quite tricky to use since any mechanical force leads to the reformation of graphite.

Our approach relied on the use of microwave-expanded graphite to form freestanding electrodes via dispersion and vacuum filtration protocols. The obtained “filter cake” is cut into round disks
of 5 mm diameter and contacted with a wire before surface modification with the enzymes,
promoter or redox mediators. This process produces interconnected porous graphite layers with
surprisingly good properties for enzyme immobilization and wiring. This work represents a
proof of concept for the use of EPG as a safe and cost-efficient alternative to other
nanostructured carbon materials for bioelectrocatalytic applications.

The aim of this proof of concept is to propose EPG as an alternative material for enzymatic
biofuel cell applications compared to commonly used CNTs [25], and graphene and its
derivatives [26]. Beside its satisfying properties for enzymatic electrocatalysis, further
advantages are, firstly, the low cost of expandable graphite (~50 € / kg), secondly, its
demonstrated non-toxicity, and thirdly, the fact that it is not a nanomaterial and therefore not
subjected to regulation.

## 2. Materials and Methods

### 2.1. Materials and Reagents.

Expandable graphite was generously provided by Mersen (France). Commercial MWCNT
proprietary blend buckypaper with reference number NTL-12218 was obtained from Nano-
TechLabs, Inc. Multi-walled carbon nanotubes (MWCNTs) (Ø=9.5 nm, purity >95%, 1.5 mm
length) were obtained from Nanocyl (Belgium) and used for lab-made buckypaper fabrication,
sodium phosphate dibasic, sodium phosphate monobasic, hexaammineruthenium(III) chloride
(98%), potassium ferricyanide(III) (99%), iron-protoporphyrin (hemin; 97%), 1,10-
phenanthroline-5,6-dione (PLQ, 97%), toluene (99.5%), and N,N-dimethylformamide (DMF,
99.9%) were purchased from Sigma-Aldrich (France). Bilirubin oxidase (BOD, 1.96 U/mg)
from *Myrothecium verrucaria* and flavin adenine dinucleotide-dependent glucose
dehydrogenase (FAD-GDH, 1,150 U/mg) from *Aspergillus* sp. were purchased from Amano
(Japan) and Sekisui Diagnostics (United Kingdom), respectively. They were stored at -20 °C
when not in use and used without further purification. Oxygen and argon were purchased from
Air Liquide (France)

## 2.2.Expanded Graphite Paper.

EPG was obtained through microwave treatment of 1 g expandable graphite in a 1 L beaker covered with a petri dish at 900 W for 30 s in a Whirlpool cook 30 microwave oven. Since this treatment provoked a tremendous sparkle formation combined with a potential evolution of toxic gases (i.e. SO₂), the experiments were performed in a fume hood or outside. The EPG volume increased by about 300 times. The finally half-filled beaker was then removed from the microwave oven and allowed to cool naturally down to room temperature. The expanded volume (EV) was calculated as the ratio of the expanded graphite volume to its mass (EV = V/m). The EPG was transferred to a graduated cylinder for volume assessment, and the average value was determined after excluding the highest and lowest volumes [27, 28].

EPG electrodes were prepared using an adjusted vacuum filtration method previously reported for fabricating lab-made CNT buckypapers (CNT-BPs) [29]. 66 mg of expanded graphite (1 mg mL⁻¹) was added to 66 mL of toluene, followed by sonication using a Bandelin Sonorex RK100 (Berlin, Germany) ultrasound bath for 90 minutes. The resulting suspension was then filtered using a diaphragm pump (MZ 2C NT model, Vacuubrand) with a Millipore PTFE filter (JHWP, with 0.45 µm pore size). The paper like EPG filter cake was rinsed with water and left to dry under vacuum for 1 hour. The EPG paper could then be removed from the filter paper and was left to dry in air overnight. The final EPG paper was then cut into individual conductive sheets with an area of 0.196 cm² (Ø = 5 mm). An electrical wire was attached to one side (the back of the electrode) with carbon paste and the sides were coated with a silicon layer to create the final EPG electrode.
Enzyme-functionalization was achieved using a drop-coating method. For the bioanode, the EPG electrode was first modified with the redox mediator. A 20 µL aliquot of a 10 mmol L⁻¹ PLQ solution in DMF was drop-coated on the EPG electrode surface and allowed to dry in air in a fume hood at room temperature. Subsequently, 20 µL of a 5 mg mL⁻¹ FAD-GDH solution in phosphate buffer (0.1 mol L⁻¹, pH 7) was spread onto the electrode surface, and the enzyme was allowed to adsorb overnight at 4°C. For the biocathode, the EPG electrode was initially functionalized with hemin to promote favorable adsorption and orientation of BOD onto the electrode surface [30]. A 20 µL aliquot of a 10 mmol L⁻¹ hemin solution in DMF was drop-coated onto the EPG electrode surface and left to dry in air in a fume hood at room temperature. After drying, 20 µL of a 5 mg mL⁻¹ BOD solution in phosphate buffer (0.1 mol L⁻¹, pH 7) was drop-cast onto the electrode surface, and the enzyme was allowed to adsorb overnight at 4°C. For comparison, CNT-BP and commercial CNT-BP electrodes were formed and modified under identical conditions and using identical quantities [31].

### 2.3. Microscopy.

Scanning electron microscopic images of the materials were obtained using a FEI/Quanta FEG 250 scanning electron microscope (SEM, Hillsboro, OR, USA) at an accelerating voltage of 5 kV.

### 2.4. Morphology

Root mean square roughness (rms) and thickness of the EPG paper were estimated from laser intensity images of the top view and vertical cross-section, respectively, using a 3D Laser Scanning Microscope (Keyence VK-X200). Specific surface area was determined from the desorption branch using the Brunauer-Emmett-Teller (BET) method. Non-modified EPG electrodes (total sample mass of ca. 15 mg) were first outgassed at 120 °C for 12 hours. N₂ adsorption/desorption isotherms were obtained at 77 K over a relative pressure range of 0.001 to 0.4. The specific surface area was calculated using the BET equation within the relative
pressure range of 0.05 to 0.17. Water contact angles were measured by depositing a 5 µL droplet
of water onto material surfaces at room temperature using a Dataphysics OCA 35 system.

2.5.Raman Spectroscopy.

Raman spectra were acquired using a WITec alpha300 RA system, which was equipped with a He/Ne laser (λ = 633 nm) and a Zeiss 50X objective. The data were then normalized to the highest peak intensity.

2.6.Electrochemical Measurements.

Electrochemical measurements were performed using a PGSTAT 100 potentiostat operated by
Nova 2.1.4 software in a conventional three-electrode electrochemical cell with an aqueous
solution (0.1 mol L⁻¹ phosphate buffer at pH 7) at a controlled temperature set at 25°C. A
platinum wire was used as the counter electrode, an Ag/AgCl (KCl saturated) as the reference
electrode, and the EPG / CNT BP electrode as the working electrode. For enzymatic biofuel
cell characterization, a two-electrode setup was used by connecting the bioanode to the counter
and reference leads, and the cathode to the working lead.

Electrochemical procedures.

2.7.Electroactive Surface Area.

The electroactive surface area (EASA) was determined using the Randles-Sevcik equation (Equation 1). Initially, the diffusion coefficients for the redox probes $Ru(NH_3)_6^{3+}/2^+$ and $Fe(CN)_6^{3-}/4^-$ (1 mmol L$^{-1}$) in phosphate buffer (0.1 mol L$^{-1}$, pH 7) were determined using a glassy carbon electrode (Ø = 3 mm). After voltammetric cycling at different scan rates from 5 mV s$^{-1}$ to 100 mV s$^{-1}$ between -0.5 V – 0.5 V ($Ru(NH_3)_6^{3+}/2^+$) and -0.4 V and 0.6 V
(Fe(CN)₆³⁻/⁴⁻), the peak currents of the redox waves were plotted versus the square root of the scan rate. The estimated diffusion coefficients for Fe(CN)₆³⁻/⁴⁻ and [Ru(NH₃)₆]³⁺/²⁺ were 1.36 × 10⁻⁶ cm² s⁻¹ and 1.91 × 10⁻⁶ cm²/s, respectively. These values were then used to estimate the EASA of the EPG electrodes and CNT-BPs examined under the same conditions.

$$I_p = (2.69 \times 10^5) n^{3/2} AD^{1/2} C\nu^{1/2} \quad \text{Equation 1}$$

$I_p$ (A) is the peak current, n is the number of electrons transferred in the redox reaction, A (cm²) is the geometric area of the electrode, C (mol cm⁻³) is the concentration of the redox species, D (cm² s⁻¹) is the diffusion coefficient and v (V s⁻¹) is the scan rate.

### 2.7.1. Specific capacitances

The specific capacitance (Equation 2) of the EPG / CNT BP electrodes were assessed from cyclic voltammograms recorded at 1, 2, 5, 10, and 20 mV s⁻¹ in phosphate buffer (0.1 mol L⁻¹, pH 7, RT), through analysis of its double-layer capacitance and relating it to the corresponding EASA (x; m² g⁻¹).

$$C_s = Q / \Delta V \quad \text{Equation 2}$$

Q is the charge measured during the discharge step and ΔV is the electrochemical potential range operated during the discharge. The recorded cyclic voltammograms are presented in Figure S2.

### 2.7.2. Surface coverage of electroactive redox species.

Surface coverages were determined from cyclic voltammograms for PLQ and hemin functionalized EPG electrodes recorded at 20 mV s⁻¹ in phosphate buffer (0.1 mol L⁻¹, pH 7, RT) by integrating the faradaic redox peaks. The surface coverage (Γ) was calculated using the Equation 3.

$$\Gamma = Q/nFA \quad \text{Equation 3}$$
Q (C) represents the integrated charge, F (96485 C mol⁻¹) is the Faraday constant, A (cm²) is the electrode's geometric area, and n denotes the number of transferred electrons.

### 2.7.3. Enzymatic fuel cell.

A bioanode and biocathode were placed face-to-face with a constant distance of 0.5 cm and immersed in phosphate buffer (0.1 mol L⁻¹, pH 7) containing 0.1 mol L⁻¹ glucose under O₂-saturated conditions (≈ 1.1 mmol L⁻¹). Polarization curves were recorded using potentiostatic discharge, where potentials ranging from 0.59 V to 0.36 V were controlled, and the resulting currents were measured to plot the enzymatic fuel cell power curve.

## 3. Results and Discussion

### 3.1. Physicochemical Characterization

Morphology and microstructure of EPG were characterized after microwave-assisted exfoliation of expandable graphite flakes. Figure 1A inset presents the SEM images of as-received expandable graphite flakes. It can be seen that this is a quite polydisperse material with particles of several hundreds of µm in diameter and a few tenths of µm in thickness. By zooming in the side of one flake (Figure 1A), a laminar composition can be observed which was likely caused by the intercalating compounds. The SEM image in Figure 1B shows the separated graphite sheets after expansion at higher magnification. The thin walls are composed of several graphene layers. No individual graphene sheet could be observed. At low magnification (inset), worm like structures up to several millimeters in length are observed. The expansion process thus led to a more than 300-fold thickness increase of the particles. It should be noted that such formed EPG is relatively fragile and mechanical pressure immediately provokes collapse to form shiny graphite sheets. Therefore, EPG was transferred to a toluene-containing round bottomed flask and then sonicated to reduce and homogenize the graphite particle sizes. After filtration and drying, free standing EPG papers were obtained. On the top side (Figure 1C) the
paper surface reveals randomly assembled graphite sheets of different sizes. The overall surface
morphology (inset) appears to be relatively rough. The section of the EPG paper (Figure 1D)
shows randomly oriented graphite with cavities or pores of several tenths of micrometers. The
sonication and filtration procedure just slightly affected the morphology of EPG.

The expanded volume, typically calculated as the ratio of the expanded graphite volume to its
mass, was estimated for the EPG to be 331 ± 12 mL g⁻¹. N₂ adsorption–desorption isotherms
revealed a BET surface area of 26 m² g⁻¹ for EPG powder. These values are comparable to those
of EPG prepared through microwave-assisted exfoliation [27, 28]. After sonication and
filtration, no substantial loss in specific surface area was observed. N₂ adsorption–desorption
isotherms revealed a BET surface area of 21 m² g⁻¹. This indicates that the ultrasonic dispersion
process, which breaks down the structure, followed by the packing of graphite sheets through
vacuum filtration, does not lead to the reformation of graphite or a significant loss of specific
surface area. Although its specific surface area is comparable to that of commercial CNTs BP
(~ 30 m² g⁻¹) [31], it is 11 times smaller than that determined for lab-made CNT BPs (~ 242 m²
g⁻¹).
<figure><img src="image_2.png" /><figcaption>**Figure 1.** SEM images for A) Expandable graphite flakes; B) Expanded graphite (EPG); and C) top view of EPG after filtration, and D) side view of the EPG paper section. All insets are the respective low magnified images.</figcaption></figure>

To gain further insight into the morphology of the EPG papers, the average surface roughness was determined using a confocal laser microscope. Figure 2A presents a top-view laser image of the EPG papers. The surface texture profile reveals the morphology of graphite sheets of ~10 µm size, and the root mean square roughness of the surface, as estimated from laser intensity images (126 x 94 µm), of 21.6 µm. This is significantly higher than the average roughness values reported for lab-made (1.9 µm) and commercial (4.8 µm) CNT-BPs [31]. Supplementary laser intensity images (not shown) of the EPG paper cross-sections revealed a thickness of 0.80 ± 0.08 mm, which is approximately three times more than lab-made CNT-BPs (0.27 ± 0.03 mm) prepared under identical conditions (quantities and treatments). Hence, the mass density of the EPG (0.10 ± 0.05 g cm⁻³) is roughly one-third to a half of the lab-made CNT-BP (0.29 ± 0.03 g cm⁻³).
Contact angle measurements (Figure 2A inset) revealed rapid water penetration into the EPG paper despite the hydrophobic character of graphite. This behavior can be explained on the one hand by a capillary effect due to the high roughness of EPG paper surface, and on the other hand, by the formation of oxygenated functions at the edges during microwave treatment, thus improving its wettability.

To assess the extent of defect sites on the graphite surfaces, Raman spectroscopy was conducted by examining the intensity ratio of the D band (disordered sp² carbon) near 1350 cm⁻¹ to the G band (crystalline sp² carbon) at 1580 cm⁻¹. Figure 2B shows representative Raman spectra of the expandable graphite precursor (top), microwave expanded graphite (middle), and the paper-shaped expanded graphite after sonication and filtration at two selected sites (bottom).

For all samples, the shape and intensity of the 2D line at 2680 cm⁻¹ clearly indicates the presence of graphite, and no traces of graphene. In some cases, surprisingly considering the violent expansion by the microwave treatment producing lot of sparkles, expandable graphite appears to have the highest amount of sp³ defects with a D/G ratio of 0.51. The D/G ratio of EPG (0.12) is quite significantly lower despite this treatment. This observation might be explained by the combustion of amorphous impurities and the re-graphitization of the basal planes at such high temperatures. The borders of such EPG are considered to be highly defective. The dispersion and filtration procedure to form the EPG paper might further have a purification effect since a D/G ratio of only 0.09 could be determined. The D/G values were determined for each sample by averaging the values of three spectra recorded at randomly chosen areas of each sample. To emphasize the high graphitization, the spectra recorded with the highest D-line intensity is presented at the bottom of Figure 2B with an acceptable D/G ratio of 0.19. Such highly graphitized surfaces are in contradiction to the contact angle results where an excellent wettability is observed. Considering the high roughness of the surface and average diameters of the EPG particles in the mm² range, and the thin walls of few graphene sheets, it can be
suggested that the defective edges are barely visible in the individual spectra. These results are
in line with previous studies found in literature [32].

To visualize our hypothesis about well-graphitized planes and defective borders, Raman maps
were recorded over several µm². Figure 2C shows the microscopic image of the mapped zone
while Figure 2D shows the 2D map of the integrated D-line intensity. Some bright zones with
high D-line intensities correspond to some of the borders of EPG particles in the microscopy
image. By superposing these two images, these features are better visualized (Figure 2E).
Diffraction or focus issues might be the reason for the differences between the microscopy
image and the Raman map.

<figure><img src="image_3.png" /><figcaption>**Figure 2:** A) Laser intensity image of the top view of the EPG paper surface (inset: water contact angle image of a 5 µL water droplet on the EPG paper surface and a section of its height</figcaption></figure>
profile) B) Raman spectra of expandable graphite (black), EPG (blue), and EPG paper (red and green) in the region between 400 and 3000 cm⁻¹. C) Microscopy image of the mapped EPG paper area, D) 2D map of the integrated D-line at 1340 cm⁻¹, and E) superposition of C and D.

### 3.2. Electrochemical Characterization

The electrochemical behavior of the expanded graphite paper electrode (EPGE) and CNT BPs, made from individual conductive sheets (Ø = 5 mm), was evaluated using two different redox probes, ruthenium hexamine ($Ru(NH_3)_6^{3+/2+}$) and hexacyanoferrate ($Fe(CN)_6^{3-/4-}$) as shown in Figure 3. $Ru(NH_3)_6^{3+/2+}$ is an outer-sphere redox probe i.e. no bond formation during electron transfer with the electrode, and is typically insensitive to electrode surface chemistry. The cyclic voltammograms, recorded at 20 mV s⁻¹ using EPGEs, exhibited well-defined cathodic and anodic peaks with mid-point potentials ($E_m$) of -0.13 V and peak-to-peak potential separations ($ΔE_p$) of 0.19 V. These values are respectively less negative and smaller compared to those of lab-made CNT-BP ($E_m$ = -0.23 V; $ΔE_p$ = 0.23 V). The more positive $E_m$ suggests that the ruthenium redox process occurs with lower overpotential on the EPGEs, while the reduced $ΔE_p$ indicates faster electron transfer kinetics or lower resistance compared to lab made CNT-BPs.

Concerning $Fe(CN)_6^{3-/4-}$, a classic redox probe whose behavior can be influenced by electrode surface properties, the EPGEs exhibited well-defined cathodic and anodic peaks with an $E_m$ of 0.24 V and a $ΔE_p$ of 0.21 V. In comparison, lab made CNT-BPs showed an $E_m$ of 0.21 V and a $ΔE_p$ of 0.22 V. The more positive $E_m$ for the EPGEs indicates that a slightly higher overpotential is required for the $Fe(CN)_6^{3-/4-}$ redox process. However, the marginal difference in $ΔE_p$ suggests that the electron transfer kinetics are similar to those observed with the lab-made CNT BPs.

Both redox probes were used to estimate the electroactive surface area (EASA) of the EPGEs. Cyclic voltammograms revealed an EASA of 3.02 cm² for $Ru(NH_3)_6^{3+/2+}$ and 3.28 cm² for $Fe(CN)_6^{3-/4-}$. Although these values are approximately half of those obtained for lab-made CNTs BPs (5.26 cm² for $Ru(NH_3)_6^{3+/2+}$ and 5.73 cm² for $Fe(CN)_6^{3-/4-}$), they are in the same
order of magnitude. These results suggest that EPGEs provide open microstructures that enable diffusion in depth of the aqueous electrolytic media, despite the clearly lower BET surface area and the high graphitization. In contrast, although lab-made CNT BPs demonstrate a higher ECSA, they are less permeable for aqueous solutions leading to a clearly reduced electroactive surface area compared to its BET surface.

Furthermore, the cyclic voltammograms reveal an approximately 14-fold lower capacitance for EPGEs (0.67 ± 0.05 F g⁻¹; 30 ± 2 µF m²) compared to lab-made CNT BPs (9.68 ± 1.09 F g⁻¹; 41.8 ± 4.5 mF m²).

<table><caption>**Table 1.** Overview of the main characteristics of the EPG paper compared to lab made CNT-BP.</caption><thead><tr><th colspan="5">Physicochemical</th><th colspan="3">Electrochemical</th></tr><tr><th></th><th>Mass Density<br>(g cm<sup>-3</sup>)</th><th>BET<br>Surface<br>(m<sup>2</sup> g<sup>-1</sup>)</th><th>Surface<br>Roughness<br>(µm)</th><th>Water<br>Contact<br>Angle<br>(°)</th><th>Electroactive Surface<br>Area (cm<sup>2</sup>)<br>Ru(NH<sub>3</sub>)<sub>6</sub><sup>3+/2+</sup> Fe(CN)<sub>6</sub><sup>3-/4-</sup></th><th>Specific<br>Capacitance<br>(F g<sup>-1</sup>)</th></tr></thead><tbody><tr><td>EPG</td><td>0.10 ± 0.05</td><td>21</td><td>21.6</td><td>Immediately<br>absorbed</td><td>3.02</td><td>3.28</td><td>0.67 ± 0.05</td></tr><tr><td>CNT BP</td><td>0.29 ± 0.03</td><td>242</td><td>1.9</td><td>56.6</td><td>5.26</td><td>5.73</td><td>9.68 ± 1.09</td></tr></tbody></table>
<figure><img src="image_4.png" /><figcaption>**Figure 3.** Cyclic voltammograms recorded at 20 mV s⁻¹ for A) EPGEs and B) CNT-BPs in Ru(NH₃)₆³⁺/²⁺ (red line) and Fe(CN)₆²⁻/₄⁻ (black line) each at a concentration of 1 mmol L⁻¹ in phosphate buffer (0.1 mol L⁻¹, pH 7, RT).</figcaption></figure>

### 3.3. Bioelectro catalytic Activity - Enzymatic Biofuel Cells

With the aim to evaluate EPG as an alternative electrode material for bioelectrocatalysis, EPGEs and CNT BPs were biofunctionalized in the same way. For bioanodes using FAD-GDH as a glucose oxidizing biocatalyst, efficient mediated electron transfer can be achieved with PLQ as the electron shuttle [29]. FAD-GDH is a favorable alternative to glucose oxidase for the bioelectrocatalytic oxidation of glucose due to its superior specificity, activity, and stability [33]. Figure 4A displays the voltammograms of the PLQ functionalized EPGEs recorded under argon. A well-defined electrochemical process with E_m at -0.10 V and a ΔE_p of 0.17 V can be observed, indicating that PLQ is accessible to the electrolyte and exhibits fast electron transfer. By varying the scan rate, the current intensity changes linearly, indicating that PLQ is immobilized on the EPG electrode (Figure 4A inset). The estimated surface coverage, Γ<sub>PLQ</sub> = 6.54 × 10<sup>-8</sup> mol cm<sup>-2</sup>, obtained from the integration of the faradaic charge, is very similar to that reported for lab-made CNT-BPs [29].

The PLQ-functionalized EPGEs were then modified with FAD-GDH as described before. The bioelectrocatalytic performance of the resulting EPG FAD-GDH-bioanode was then assessed
in the presence of 0.1 mol L⁻¹ glucose in phosphate buffer (0.1 mol L⁻¹, pH 7). The cyclic voltammogram (Figure 4B) shows an almost sigmoidal current profile with a clear oxidation wave arising from the PLQ. This behavior is quite typical since during the enzymatic glucose oxidation, this electron shuttle is reduced by the enzyme and then re-oxidized during the anodic scan, finalizing the mediated electron transfer. A stable electrocatalytic current is obtained beyond 0.2 V. The efficient use of the mediator during the mediated electron transfer reaction is represented by the absence of a reduction wave during the cathodic scan. At 0.3 V, the FAD-GDH-functionalized EPGE achieved a catalytic current of 2.87 mA cm⁻², which is approximately half of the maximum value obtained for lab-made CNTs BP (6.11 mA cm⁻²), as shown in Figure 4C. To gain more insight into the appropriateness of EPG for bioelectrocatalysis, the performance was compared with commercial CNT-BP bioanodes having an identical diameter, and prepared under identical conditions. These CNT-BP bioelectrodes gave an electrocatalytic current of 1.37 mA cm⁻² at 0.3 V, which is clearly less performing than the presented EPG bioanode (Figure 4D). Furthermore, we estimated the bioelectrocatalytic stability of the EPG bioanode by continuous cycling between -0.4 V and 0.4 V (Figure S1A). After 30 cycles, the catalytic current at 0.3 V decreased from 2.87 mA cm⁻² to 1.52 mA cm⁻², which represents a loss of 47.2 %. Such stability is not excellent but comparable to former studies using CNT deposits on glassy carbon electrodes [34]
<figure><img src="image_5.png" /><figcaption>Figure 4: A) Cyclic voltammograms in the range between -0.4 V and 0.4 V recorded in phosphate buffer (0.1 M, pH 7) at different scan rates from 5 to 100 mV s⁻¹ under argon using PLQ-functionalized EPGEs. Inset: plot of the current peak intensities as a function of the scan rate. B)-D) cyclic voltammograms recorded at 2 mV s⁻¹ in phosphate buffer (0.1 mol L⁻¹ at pH 7) for B) FAD GDH-functionalized EPGEs, C) CNT BP, and D) commercial CNT BP in absence (black), and in presence of 0.1 mol L⁻¹ glucose (red). For B)-D), the current density axis is in the same range for better clarity.</figcaption></figure>

For the design of the EPG biocathode, bilirubin oxidase (BOD) was chosen due to its reliable and attractive electrocatalytic performances [35]. This biocatalyst belongs to the multicopper enzyme family, which reduces O₂ to H₂O in nature by oxidizing bilirubin, a metabolite of hemin. By incorporating hemin molecules at electrodes, the porphyrin structure allows stronger π stacking interactions with graphitized carbon while the bilirubin ensures a strong affinity with
the BOD, in part due to the carboxylate functionalities. BOD is not capable to oxidize hemin,
leading to an orientated immobilization that brings one catalytic copper center close to the
electrode surface, thus allowing direct electron transfer [30].

After drop-coating hemin on the EPGEs, cyclic voltammograms were recorded at different scan
rates. Figure 5A shows voltammograms obtained for the hemin-functionalized EPGE recorded
under argon. The well-defined Fe²⁺/³⁺ redox couple signal at E_m = -0.33 V and an almost perfect
linearity of the peak current intensities towards the scan rate (R² = 0.995, Figure 5A inset)
demonstrates the firm adsorption of hemin on EPG. The surface coverage of Γ_hemin = 4.84 ×
10⁻⁸ closely matches the values previously reported by our research group for hemin-modified
lab-made CNT-BPs [36]. Figure 5B shows the cyclic voltammograms recorded in Ar and in
O₂-saturated buffer solutions for the BOD-functionalized EPGE where a catalytic current of -
0.41 mA cm⁻² at 0.2 V could be obtained. This bioelectroreduction performance remains
comparable to the values obtained for lab-made CNT-BPs (-1.30 mA cm⁻²) under identical
conditions (Figure 5C). Using commercial CNT-BPs, the performance is just slightly superior
to EPG biocathodes with -0.58 mA cm⁻² at 0.2 V. However, parts of this current are of
capacitive nature, which lowers this superiority. Artificial aging by cycling in the range between
0 V and 0.7 V, the catalytic current at 0.2 V decreased by 45.8% after 30 continuous cycles
(from 0.41 mA cm⁻² to -0.22 mA cm⁻²) as shown in Figure S1B.

It should be noted that the catalytic current densities were calculated as commonly practiced
using the geometric electrode surface (Ø = 5 mm, 0.196 cm²). Considering the determined
EASAs of EPGE and lab-made CNT-BPs, comparable bioelectro catalytic activities were
observed. EPG bioanodes and biocathodes provided respective current densities of 179 µA cm⁻²
and -26.6 µA cm⁻², while lab-made CNT-BPs provided 201 µA cm⁻² and -44.5 µA cm⁻²,
respectively. Furthermore, when considering the BET surface area and not the geometric area,
EPG presents a significant advantage over lab-made CNT-BPs. EPG bioelectrodes reach anodic
and cathodic current densities of 25.84 µA cm⁻² and -3.73 µA cm⁻², respectively, whereas lab-made CNT-BPs yield 4.77 µA cm⁻² and -1.05 µA cm⁻². It seems to be clear that most of the specific surface area of lab-made CNT-BPs remains inaccessible for the electrolyte and biofunctionalizations. Lab-made CNT-BPs are typically composed of densely packed nanometric CNTs forming primarily micro-mesoporous structures. Despite the small size of enzymes, they may have difficulty to penetrate in the pores and permeate deep inside or through the entwined CNTs network. For instance, we recently described the assembly of two CNT-BP sheets forming a microcavity that effectively traps enzymes without leaking [36]. In contrast, the EPG is composed of micrometric graphite sheets arranged as interconnected porous microsheets, which provides a graphitic open pore system that seems to facilitate enzyme penetration, permeation, and adsorption.

<figure><img src="image_6.png" /><figcaption>**Figure 5:** A) Cyclic voltammograms in the range between -0.8 V and 0.2 V recorded in</figcaption></figure>
phosphate buffer (0.1 mol L⁻¹, pH 7) at different scan rates from 5 to 100 mV s⁻¹ under argon using Hemin-functionalized EPGEs. Inset: plot of the current peak intensities as a function of the scan rate. B)-D) cyclic voltammograms in the range between 0 and 0.7 V recorded at 2 mV s⁻¹ in phosphate buffer (0.1 mol L⁻¹ at pH 7) for B) BOD-functionalized EPGEs, C) CNT BP, and D) commercial CNT BP in Ar saturated (black), and O₂ saturated (red) solution. For B)-D), the current density axis is in the same range for better clarity.

It should be noted that the catalytic current densities were calculated as commonly practiced using the geometric electrode surface (Ø = 5 mm, 0.196 cm²). Considering the determined EASAs of EPGE and lab-made CNTs buckypapers, comparable bioelectrocatalytic activities were observed. EPG bioanodes and biocathodes provided respective current densities of 179 µA cm⁻² and -26.6 µA cm⁻², while lab-made CNT-BPs provided 201 µA cm⁻² and -44.5 µA cm⁻², respectively. Furthermore, when considering the BET surface area and not the geometric area, EPG presents a significant advantage over lab-made CNT-BPs. EPG bioelectrodes reach anodic and cathodic current densities of 25.84 µA cm⁻² and -3.73 µA cm⁻², respectively, whereas lab-made CNTs BPs yield 4.77 µA cm⁻² and -1.05 µA cm⁻². It seems to be clear that most of the specific surface area of lab-made CNT-BPs remains inaccessible for the electrolyte and biofunctionalizations. Lab-made CNT-BPs are typically composed of densely packed nanometric CNTs forming primarily micro-mesoporous structures. Despite the small size of enzymes, they may have difficulty to penetrate in the pores and permeate deep inside or through the entwined CNTs network. For instance, we recently described the assembly of two CNT-BP sheets forming a microcavity that effectively traps enzymes without leaking [36]. In contrast, the EPG is composed of micrometric graphite sheets arranged as interconnected porous microsheets, which provides a graphitic open pore system that seems to facilitate enzyme penetration, and permeation. However, the main issue, which has to be addressed in future work, is the unsatisfying repeatability for the fabrication of EPG bioelectrodes related to their
fragile morphology. Each manipulation risks even slight compressions of the material, which provokes drastic changes in performances. We could nonetheless form two bioanodes and two biocathodes providing similar, and up to date optimal electrocatalytic behavior with 3.38 mA cm⁻² and 2.87 mA cm⁻² for the bioanodes at 0.3 V (Figure S3A, B) and 0.41 mA cm⁻² and 0.32 mA cm⁻² for the biocathodes at 0.2 V (Figure S3 C, D).

After demonstrating promising bioelectrocatalytic activity, FAD-GDH- and BOD-functionalized EPGEs were set face-to-face in a classical biofuel cell setup. The enzymatic EPG glucose biofuel cell exhibited a notable OCV of 0.65 V which matches perfectly with the OCV of our former work using lab-made CNT-BPs under quiescent and O₂ saturated conditions [29]. Potentiostatic discharge experiments (Figure 6A) revealed a maximum current density of 0.37 mA cm⁻² and a peak power output of 0.15 mW cm⁻² at 0.41 V (Figure 6B) by measuring the current after 10 s discharge of the capacitive current. This performance is again below the power output of lab-made CNT-BPs, but these results are nicely in line with the electroactive surface area of EPG. Nonetheless, compared to glucose biofuel cells using other nanostructures carbon materials, these performances are comparable [6].

<figure><img src="image_7.png" /><figcaption>**Figure 6.** A) Potentiostatic discharge curves recorded from 0.59 V to 0.36 V for 10 s in 0.1 mol L⁻¹ glucose under O₂-saturated conditions (≈ 1.1 mmol L⁻¹) B). Polarization (black cubes) and</figcaption></figure>
power density (red cubes) curves for enzyme-functionalized EPG paper fuel cell from the potentiostatic discharges.

4. Conclusion

This work is considered as a proof of concept to permit the use of a cheap and non-toxic alternative to nanostructured carbon materials for enzymatic bioelectrocatalysis. The catalytic and energy conversion performances are at this stage still lower, but there is still room for optimization. EPG bioelectrodes can be fabricated easily and were successfully applied in bioelectrocatalytic oxidation and reduction reactions for the enzymatic biofuel cell application. EPG obtained by microwave-assisted expansion of expandable graphite flakes was re-dispersed and filtered to obtain a relatively stable paper from which freestanding paper-type electrodes could be formed. The processed expanded graphite exhibited promising physicochemical properties, including well-packed graphite sheets with low mass density, high surface roughness, wettability, and a relatively high specific surface area for a microsized material. As an electrode, the processed expanded graphite displayed electron transfer reactivity comparable to that of carbon nanotube electrodes. The microstructure formed by the graphite sheets creates open pathways that facilitate efficient diffusion of diffusional probe chemicals and enhance biocatalyst penetration. Bioelectrocatalytic studies confirmed the ability to support effectively electroenzymatic oxidation-reduction processes through both direct and mediated electron transfer. As an enzymatic biofuel cell, the power output was lower but comparable to other carbon-based systems. Regarding the material costs, expandable graphite is three orders of magnitude cheaper than thin multiwalled carbon nanotubes. By simply increasing the amounts of EPG used in electrode systems, the performances should be equalized. However, mechanical issues, which cause unsatisfying reproducibility, should be addressed. Stability issues could potentially be overcome with the incorporation of additives such as conductive polymers. This
innovative approach to use expanded graphite could pave the way for a new generation of cost-efficient carbon-based electrodes.

**Table of abbreviations:**

<table><tr><td>9,10 phenanthroline-5,6-dione</td><td>PLQ</td></tr><tr><td>bilirubin oxidase</td><td>BOD</td></tr><tr><td>Brunauer-Emmett-Teller</td><td>BET</td></tr><tr><td>carbon nanotube buckypaper</td><td>CNT-BP</td></tr><tr><td>carbon nanotubes</td><td>CNTs</td></tr><tr><td>cyclic voltammogram</td><td>CV</td></tr><tr><td>dimethylformamide</td><td>DMF</td></tr><tr><td>direct electron transfer</td><td>DET</td></tr><tr><td>electroactive surface area</td><td>EASA</td></tr><tr><td>expanded graphite paper electrode</td><td>EPGE</td></tr><tr><td>expanded popcorn graphite</td><td>EPG</td></tr><tr><td>expanded volume</td><td>EV</td></tr><tr><td>FAD- glucose dehydrogenase</td><td>FAD-GDH</td></tr><tr><td>mediated electron transfer</td><td>MET</td></tr><tr><td>mid-point potential</td><td>Em</td></tr><tr><td>multi-walled carbon nanotubes</td><td>MWCNTs</td></tr><tr><td>open circuit voltage</td><td>OCV</td></tr><tr><td>peak-to-peak potential separation</td><td>Ep</td></tr><tr><td>root mean square roughness</td><td>rms</td></tr><tr><td>scanning electron microscopy</td><td>SEM</td></tr></table>

**Declaration of interests**

A.M, A.J.G and M.H. hold a patent registration as inventors related to the technology described in this paper. The patent is registered under the following reference:

International Application No.: PCT/EP2024/069105

Title: Expanded ultraporous graphite electrodes for (bio)electronics device
Applicants: Centre National de la Recherche Scientifique; Université Grenoble Alpes.

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

Author contributions

The manuscript was written through contributions of all authors. Conceptualization: P.H.M.B., A.M., and M.H.; Methodology: P.H.M.B., A.M., A.J.G. Y.N. and M.H.; formal analysis: P.H.M.B., A.M., A.J.G. P.A., Y.N. and M.H.; investigation: P.H.M.B., A.M. A.J.G. and M.H.; data curation: P.H.M.B. A.M. and M.H.; validation: P.H.M.B. A.J.G. and M.H.; writing - original draft preparation: P.H.M.B. and M.H.; writing - review and editing: P.H.M.B, A.J.G. M.H.; supervision: M.H.; funding acquisition: M.H. All authors have given approval to the final version of the manuscript

Acknowledgments

The authors wish to acknowledge the support from the platform Chimie NanoBio ICMG FR 2607 (PCN-ICMG), from the LabEx ARCANE (ANR-11-LABX-0003-01 and CBH-EUR-GS, ANR-17-EURE-0003), and.an from the Institut Carnot PolyNat (CARN 0007-01). The authors particularly thank the project Superbiocap (ANR-21-CEOS-0020·02) for financial support.

References

[1] Evgeny Katz, Paolo Bollella, Fuel Cells and Biofuel Cells: From Past to Perspectives, Isr. J. Chem. 60 (2020) 1-18.

[2] V.M. Reports®, Global Enzymatic Biofuel Cell Market By Type (Direct Electron Transfer (DET), Mediator Electron Transfer (MET)), By Application (Wearable Consumer Electronics, Implantable Medical Devices), By Geographic Scope And Forecast, (2024), Report ID: 595318
Study Period : 2021-2031; https://www.verifiedmarketreports.com/product/enzymatic-biofuel-cell-market/

[3] Caroline Abreu, Yannig Nedellec, Olivier Ondel, Francois Buret, Serge Cosnier, Alan Le Goff, Michael Holzinger, Towards eco-friendly power sources: In series connected glucose biofuel cells power a disposable ovulation test, Sens. Actuator B-Chem. 277 (2018) 360-364.

[4] Alexandre Cheikhi, Meet BeFC, the scientists replacing lithium batteries with compostable paper strips 2025. https://newenergy-today.com/news/meet-befc-the-scientists-replacing-lithium-batteries-with-compostable-paper-strips/.

[5] Ross D. Milton, David P. Hickey, Sofiene Abdellaoui, Koun Lim, Fei Wu, Boxuan Tan, Shelley D. Minteer, Rational design of quinones for high power density biofuel cells, Chem. Sci. 6(8) (2015) 4867-4875.

[6] Sufia ul Haque, Abu Nasar, Narcis Duteanu, Sadanand Pandey, Inamuddin, Carbon based-nanomaterials used in biofuel cells – A review, Fuel 331 (2023) 125634.

[7] A. J. Gross, M. Holzinger, S. Cosnier, Buckypaper bioelectrodes: emerging materials for implantable and wearable biofuel cells, Energy Environ. Sci. 11(7) (2018) 1670-1687.

[8] Michael Holzinger, Raoudha Haddad, Alan Le Goff, Serge Cosnier, Enzymatic Glucose Biofuel Cells: Shapes and Growth of Carbon Nanotube Matrices, in: S.E. Lyshevski (Ed.), Dekker Encyclopedia of Nanoscience and Nanotechnology, Third Edition, CRC Press2016, pp. 1-10.

[9] A. B. Kane, R. H. Hurt, H. Gao, The asbestos-carbon nanotube analogy: An update, Toxicol Appl Pharmacol 361 (2018) 68-80.

[10] M. R. Chetyrkina, F. S. Fedorov, A. G. Nasibulin, In vitro toxicity of carbon nanotubes: a systematic review, RSC Adv 12(25) (2022) 16235-16256.
[11] Shoji Fukushima, Tatsuya Kasai, Yumi Umeda, Makoto Ohnishi, Toshiaki Sasaki, Michiharu Matsumoto, Carcinogenicity of multi-walled carbon nanotubes: challenging issue on hazard assessment, J Occup Health 60(1) (2018) 10-30.

[12] J. A. Chavez-Hernandez, A. J. Velarde-Salcedo, G. Navarro-Tovar, C. Gonzalez, Safe nanomaterials: from their use, application, and disposal to regulations, Nanoscale Adv 6(6) (2024) 1583-1610.

[13] European Chemicals Agency (ECHA), Nanomaterials, 2020. https://echa.europa.eu/regulations/nanomaterials.

[14] B. Brodie, Note sur un nouveau procédé pour la purification et la désagrégation du graphite, Ann. Chem. Physique 45 (1855) 351.

[15] Ulrich Hofmann, Walter Rüdorff, The formation of salts from graphite by strong acids, Transactions of the Faraday Society 34 (1938) 1017-1021.

[16] M. S. Dresselhaus, G. Dresselhaus, Intercalation compounds of graphite, Advances in Physics 51(1) (2002) 1-186.

[17] Meng Lu Wang, Li Ji, Expansion Mechanism of Expandable Graphite Formed by Natural Graphite with Different Particle Size, Advanced Materials Research 499 (2012) 16-19.

[18] Haiying Lu, Xianghong Chen, Jiakui Zhang, Yu Lei, Wenlu Min, Jiantie Xu, Graphite and its main applications, Functional Carbon Materials, IOP Publishing, 2022, pp. 6-1-6-28.

[19] Bartłomiej Mazela, Anyelkis Batista, Wojciech Grześkowiak, Expandable Graphite as a Fire Retardant for Cellulosic Materials—A Review, Forests 11(7) (2020) 755.

[20] P. Murugan, R. D. Nagarajan, B. H. Shetty, M. Govindasamy, A. K. Sundramoorthy, Recent trends in the applications of thermally expanded graphite for energy storage and sensors - a review, Nanoscale Adv 3(22) (2021) 6294-6309.
[21] X. K. Yu, Y. B. Tao, Improvement of thermal cycle stability of paraffin/expanded graphite composite phase change materials and its application in thermal management, Journal of Energy Storage 63 (2023) 107019.

[22] S. Eigler, Graphite sulphate - a precursor to graphene, Chem Commun (Camb) 51(15) (2015) 3162-5.

[23] Jerome Rajendran, Brahmari H. Shetty, Dhanraj Ganapathy, Preethika Murugan, Raji Atchudan, Dhamodharan Umapathy, Ajit Khosla, Ashok K. Sundramoorthy, Thermally Expanded Graphite Incorporated with PEDOT:PSS Based Anode for Microbial Fuel Cells with High Bioelectricity Production, Journal of The Electrochemical Society 169(1) (2022) 017515.

[24] Luye Chen, Youzhi Li, Jiani Yao, Gaoming Wu, Bin Yang, Lecheng Lei, Yang Hou, Zhongjian Li, Fast expansion of graphite into superior three-dimensional anode for microbial fuel cells, Journal of Power Sources 412 (2019) 86-92.

[25] Michael Holzinger, Functionalization of Carbon Nanotubes for Bioelectrochemical Applications, IEEE Nanotechnology Materials and Devices Conference (NMDC) IEEE Xplore, Paestum, Italy, 2023, pp. 115-119.

[26] J. Tang, X. Yan, C. Engelbrekt, J. Ulstrup, E. Magner, X. Xiao, J. Zhang, Development of graphene-based enzymatic biofuel cells: A minireview, Bioelectrochemistry 134 (2020) 107537.

[27] Ting Liu, Ruijun Zhang, Xuesha Zhang, Kang Liu, Yanyan Liu, Pengtao Yan, One-step room-temperature preparation of expanded graphite, Carbon 119 (2017) 544-547.

[28] Qun Wei, Lei Xu, Zhimeng Tang, Zhangbiao Xu, Cheng Xie, Lirong Guo, Wei Li, High-performance expanded graphite from flake graphite by microwave-assisted chemical intercalation process, Journal of Industrial and Engineering Chemistry 122 (2023) 562-572.
[29] Andrew J. Gross, Xiaohong Chen, Fabien Giroud, Caroline Abreu, Alan Le Goff, Michael Holzinger, Serge Cosnier, A High Power Buckypaper Biofuel Cell: Exploiting 1,10-Phenanthroline-5,6-dione with FAD-Dependent Dehydrogenase for Catalytically-Powerful Glucose Oxidation, ACS Catal. 7 (2017) 4408-4416.

[30] Noémie Lalaoui, Alan Le Goff, Michael Holzinger, Serge Cosnier, Fully Oriented Bilirubin Oxidase on Porphyrin-Functionalized Carbon Nanotube Electrodes for Electrocatalytic Oxygen Reduction, Chem. Eur. J. 21(47) (2015) 16868-16873.

[31] Xiaohong Chen, Andrew J. Gross, Fabien Giroud, Michael Holzinger, Serge Cosnier, Comparison of Commercial and Lab-made MWCNT Buckypaper: Physicochemical Properties and Bioelectrocatalytic $O_2$ Reduction, Electroanalysis 30(7) (2018) 1511-1520.

[32] Walter Wilhelm Focke, Heinrich Badenhorst, Washington Mhike, Hermanus Joachim Kruger, Dewan Lombaard, Characterization of commercial expandable graphite fire retardants, Thermochimica Acta 584 (2014) 8-16.

[33] Ross D. Milton, Fabien Giroud, Alfred E. Thumser, Shelley D. Minteer, Robert C. T. Slade, Hydrogen peroxide produced by glucose oxidase affects the performance of laccase cathodes in glucose/oxygen fuel cells: FAD-dependent glucose dehydrogenase as a replacement, Physical Chemistry Chemical Physics 15(44) (2013) 19371-19379.

[34] Paulo Henrique M. Buzzetti, Pierre-Yves Blanchard, Emerson Marcelo Girotto, Yuta Nishina, Serge Cosnier, Alan Le Goff, Michael Holzinger, Insights into carbon nanotube-assisted electro-oxidation of polycyclic aromatic hydrocarbons for mediated bioelectrocatalysis, Chemical Communications 57 (2021) 8957-8960.

[35] B. Thangavel, G. Venkatachalam, J. H. Shin, Emerging Trends of Bilirubin Oxidases at the Bioelectrochemical Interface: Paving the Way for Self-Powered Electrochemical Devices and Biosensors, ACS Appl. Bio Mater 7(3) (2024) 1381-1399.
[36] Anastasiia Berezovska, Paulo Henrique M. Buzzetti, Yannig Nedellec, Chantal Gondran, Fabien Giroud, Andrew J. Gross, Stephane Marinesco, Serge Cosnier, A hollow microcavity enzymatic fuel cell for in vivo energy harvesting, Cell Reports Physical Science 5(9) (2024) 102203.