Biosensors and Bioelectronics 42 (2013) 219–224
Contents lists available at SciVerse ScienceDirect
Biosensors and Bioelectronics
journal homepage: www.elsevier.com/locate/bios
Direct electrochemistry and Os-polymer-mediated bioelectrocatalysis
of NADH oxidation by Escherichia coli flavohemoglobin at graphite electrodes
Maciej Sosna a, Alessandra Bonamore b, Lo Gorton c, Alberto Boffi b, Elena E. Ferapontova a,n
a
Interdisciplinary Nanoscience Center (iNANO), Aarhus University, Gustav Wieds Vej 14, DK-8000 Aarhus C, Denmark
Department of Biochemical Sciences and CNR Insitute of Molecular Biology and Pathology, University La Sapienza, 00185 Rome, Italy
c
Department of Analytical Chemistry/Biochemistry and Structural Biology, Lund University, P.O. Box 124, SE-221 00 Lund, Sweden
b
a r t i c l e i n f o
abstract
Article history:
Received 6 October 2012
Received in revised form
25 October 2012
Accepted 30 October 2012
Available online 7 November 2012
Escherichia coli flavohemoglobin (HMP), which contains one heme and one FAD as prosthetic groups
and is capable of reducing O2 by its heme at the expense of NADH oxidized at its FAD site, was
electrochemically studied at graphite (Gr) electrodes. Two signals were observed in voltammograms of
HMP adsorbed on Gr, at 477 and 171 mV vs. Ag9AgCl, at pH 7.4, correlating with electrochemical
responses from the FAD and heme domains, respectively. The electron transfer rate constant for ET
reaction between FAD of HMP and the electrode was estimated to be 83 s 1. Direct bioelectrocatalytic
oxidation of NADH by HMP was not observed, presumably due to impeded substrate access to HMP
orientated on Gr through the FAD-domain and/or partial denaturation of HMP. Bioelectrocatalysis was
achieved when HMP was wired to Gr by the Os redox polymers, with the onset of NADH oxidation at
the formal potential of the particular Os complex (þ140 mV or 195 mV). Apparent Michaelis
and jmax were determined, showing bioelectrocatalytic efficiency of NADH oxidation
constants Kapp
M
by HMP exceeding the one earlier shown with diaphorase, which makes HMP very attractive as a
component of bioanalytical and bioenergetic devices.
& 2012 Elsevier B.V. All rights reserved.
Keywords:
Escherichia coli flavohemoglobin
NADH
Os-complex redox polymers
Bioelectrocatalysis
1. Introduction
Nicotinamide adenine dinucleotide (NAD) and nicotinamide adenine dinucleotide phosphate (NADP) are pyridine coenzymes
involved in electron transport in the mitochondrial respiratory chain
(the NAD þ /NADH couple) and in the reductive biosynthesis (NADP þ /
NADPH) (Nicholls and Ferguson, 2002). Over 400 NAD(P) þ -dependent dehydrogenases use NAD(P) þ as the electron acceptor recycling
the enzyme after reaction with the substrate (Gorton and Domı́nguez,
2007; Lobo et al., 1997), and these biocatalytic reactions find wide
applications in electrochemical biosensors (Gorton and Domı́nguez,
2007; Lobo et al., 1997; Maka et al., 2003) and biofuel cell development (Minteer et al., 2007). Therewith, it is crucial that NAD(P)H
produced during the biocatalytic cycle can be electrochemically
regenerated. However, direct electrochemical oxidation of NAD(P)H
proceeds with a substantial overpotential and thus suffers from
interfering electrooxidation of other species; it is irreversible and
often leads to the electrode fouling (Moiroux and Elving, 1980).
To overcome these problems modified electrodes were extensively
used to reduce the overpotential of the reaction and to maintain fast
reaction kinetics. Modified electrodes for NADH oxidation exploit
n
Corresponding author.
E-mail address: [email protected] (E.E. Ferapontova).
0956-5663/$ - see front matter & 2012 Elsevier B.V. All rights reserved.
http://dx.doi.org/10.1016/j.bios.2012.10.094
conductive polymers and different types of mediators such as
quinones, diimines, flavins, phenothiazines and phenoxazine derivatives, also polymerized (Bartlett and Simon, 2003; Bartlett et al., 1991;
Gorton, 1986; Gorton and Dominguez, 2002, 2007; Rincon et al.,
2010; Sandström et al., 2000). Another approach involves bioelectrocatalytic oxidation of NADH, however, in practice just a few
flavoenzymes were successfully applied for direct (unmediated)
NADH oxidation (Barker et al., 2007; Kobayashi et al., 1992; Reeve
et al., 2012; Zu et al., 2003); otherwise, enzyme wiring to electrodes
by a suitable redox mediator was used (Antiochia and Gorton, 2007;
Tasca et al., 2008; Tsujimura et al., 2002a).
In the present work we electrochemically studied flavohemoglobin from Escherichia coli (HMP) and its bioelectrocatalytic
properties in the reaction of NADH oxidation. HMP is a 43 kDa
two-domain soluble protein, which contains one heme of a b-type
and one FAD as prosthetic groups, and is capable of reducing
oxygen to superoxide by its heme at the expense of NADH
oxidized at its FAD site (Ioannidis et al., 1992; MembrilloHernandez et al., 1996; Poole et al., 1994). Therewith FAD ‘‘relays’’
electron transfer (ET) from NADH to the heme and further to
the Fe(II)-heme-bound oxygen. The physiological role of HMP,
particularly under anaerobic conditions, is still not fully clear,
however, its enzymatic properties, namely the catalysis of NADH
oxidation are very attractive with respect to possible bioanalytical
and bioenergetic applications.
220
M. Sosna et al. / Biosensors and Bioelectronics 42 (2013) 219–224
Fig. 1. (A). Representative cyclic voltammograms recorded with the HMP-modified graphite electrode at potential scan rates n ranging between 20 and 350 mV s 1.
(B). Potential scan rate dependence of the voltammetric peak currents. (C). Scan rate dependence of the FAD peak potentials (Ep) vs. the mid-peak potential (Em). CVs were
recorded in deoxygenated 20 mM Tris buffer, pH 7.4.
Here, the direct electrochemistry of HMP was studied on
graphite electrodes, and the ability of HMP to directly catalyze
NADH oxidation in a mediatorless electron transfer (ET) reaction
with the electrode was explored. In the alternative reagentless
approach, electronic wiring of HMP to the electrodes by two types
of osmium complex containing polymers (Fig. 1S, SI), avoiding the
diffusion of the mediator out of the enzyme film at the electrode
surface, has been used. This wiring approach was previously used
to achieve bioelectrocatalysis of such FAD containing enzymes as
glucose oxidase (Gregg and Heller, 1990; Mao et al., 2003),
diaphorase (Antiochia and Gorton, 2007; Tasca et al., 2008;
Tsujimura et al., 2002a), pyranose oxidase (Timur et al., 2006;
Zafar et al., 2010), FAD-dependent glucose dehydrogenase (Zafar
et al., 2012) and in the development of O2-reducing biocathodes
based on blue-copper enzymes such as laccases (Barriere et al.,
2004; Daigle et al., 1998) and bilirubin oxidase (Mano et al., 2002;
Tsujimura et al., 2002b). It was also successfully used for wiring
of the redox centers of such a fragile membrane enzyme complex
as theophylline oxidase (Shipovskov and Ferapontova, 2008),
providing retention of its bioelectrocatalytic activity within the
polymer matrix (Ferapontova et al. , 2006), and even those of whole
cells (Coman et al., 2009; Hasan et al., 2012; Vostiar et al., 2004).
2. Materials and methods
2.1. Reagents
Poly(ethyleneglycol) 400 diglycidyl ether (PEGDGE) (Polysciences Inc.), Trizmas hydrochloride (Sigma), nicotinamide adenine dinucleotide, reduced form (NADH) and components of
buffer solutions (Sigma-Aldrich) were used as received. Redox
polymers:
poly
(1-vinylimidazole)12-[Os(4,40 -dimethyl-2,20 bipyridyl)2 Cl2]2 þ /3 þ (redox polymer I) and poly(vinylpyridine)[osmium-(N,N0 -methyl-2,20 -biimidazole)3]2 þ /3 þ complex (redox
polymer II) were a generous gift from TheraSense (Alameda, CA,
USA, now Abbot Diagnostics). Their synthesises were previously
described (Mao et al., 2003; Ohara et al., 1994) and formal redox
potentials were determined as þ140 mV and 195 mV vs.
Ag/AgCl (0.1 M KCl) for polymers I and II, respectively. Structures
of both polymers can be found in ESI (Fig. S1). Flavohemoglobin
from E. coli (HMP) was purified as described elsewhere (Bonamore
et al., 2001). All solutions were prepared with deionised Milli-Q
water (18.2 MO cm, Millipore, Bedford, MA, USA). When required, the
working solutions were deoxygenated with N2 (AGA, Denmark).
2.2. Electrochemical experiments
Cyclic voltammetry (CV) and differential pulse voltammetry (DPV)
experiments were performed in a single compartment glass cell using
mAutolab Type III potentiostat (Eco Chemie, The Netherlands)
equipped with a GPES 4.9 software. A standard three electrode
arrangement was used. The working electrodes were 3.05 mm
diameter rods of solid spectroscopic graphite (SGL Carbon AG, Werk
Ringsdorff, Bonn, Germany, type RW001) inserted in home-made
PTFE holders; a platinum wire was the counter electrode and Ag9AgCl
(3 M KCl) (Metrohm) was the reference electrode. All potentials in the
remainder of the article are quoted with respect to Ag9AgCl (3 M KCl).
In DPV, the modulation amplitude was 50 mV, the modulation time
was 50 ms, and interval time was 0.2 s.
2.3. Electrodes preparation
Graphite (Gr) electrodes were polished on silicon carbide
paper (grade 1000, Struers), rinsed with a copious amount of
water and dried in a stream of N2. The surface was subsequently
modified with either HMP (10 ml, 4.1 mg ml 1 HMP solution in
10 mM phosphate buffer solution, pH 7 (PBS)) or a mixture of
HMP (5 ml, 4.1 mg ml 1 HMP solution in 10 mM PBS), Os redox
polymer (1 ml, 10 mg ml 1 in water) and cross-linker PEGDGE
(1 ml, 2.5 mg ml 1 in water). The chosen preparation method was
by drop casting on the freshly polished Gr electrodes, which were
incubated at 4 1C for 5 h before subsequent washing in 20 mM
Tris buffer, pH 7.4, and electrochemical characterization.
3. Results and discussion
3.1. Direct electrochemistry of HMP adsorbed on graphite
Direct electrochemistry of HMP adsorbed on Gr electrodes
revealed a pronounced redox couple with a midpoint potential of
477 mV at pH 7.4 (Fig. 1A), correlating with an FAD/FADH2
redox transformation ( 445 mV, pH 7.45) (Gorton and
Johansson, 1980; Lowe and Clark, 1956). Another redox couple,
at 62 mV was ascribed to the redox conversion of the surface
M. Sosna et al. / Biosensors and Bioelectronics 42 (2013) 219–224
100
0
j , µA cm
-2
30
20
-100
-200
-300
-400
-1.0
-0.5
0.0
0.5
E vs Ag/AgCl , V
10
0
-0.8
-0.6
-0.4
-0.2
0.0
0.2
0.4
Fig. 2. Background corrected differential pulse voltammogram of HMP adsorbed
on graphite (thick solid line). Thin solid lines are fits of the peaks to the
experimental data, the dotted line is a sum of the fitted peaks. Inset: cyclic
voltammograms at HMP modified graphite electrode in N2 (dotted line) and air
(solid line) saturated 20 mM Tris buffer, pH 7.4; n ¼ 30 mV s 1.
quinones as it was also observed in the voltammograms in the
absence of HMP. Cyclic voltammetry did not reveal any appreciable redox peaks originating from the heme of HMP.
In DPV, however, a distinct shoulder can be seen at 171 mV
suggesting a direct electrical contact between the heme iron and
the electrode (Fig. 2). Assuming 2e and 1e transfer processes
for FAD and heme, respectively, the DPV peak ratio of 4:1 is
expected if the equivalent amounts of FAD and heme are electronically contacted with the electrode (Ferapontova et al., 2003).
The comparison of the peak currents of de-convoluted DPV peaks
yielded the FAD to heme ratio of 7:1 and is indicative of excess of
electrochemically active FAD over the heme active sites.
The amount of the FAD centers directly communicating with
the electrode surface obtained by integration of the CV peaks
(a 2e redox process) was 490710 pmol cm 2 (per geometric
surface area). This number is significantly higher than the theoretical monolayer enzyme coverage estimated on the basis of the
molecular dimensions of HMP (Ilari et al., 2002) as 3–5.5 pmol cm 2
(depending on the enzyme orientation). Considering the high
surface roughness of the Gr electrodes used (the roughness factor
above 5 (Jaegfeldt et al., 1983)), the HMP coverage corrected for the
real electrode surface area decreases to 100 pmol cm 2, which is
still unusually high. It suggests that the majority of the FAD
voltammetric signal is likely to originate from FAD released from
HMP due to protein unfolding. The potential of the FAD/FADH2
redox couple seen here is identical to that of free adsorbed FAD on
graphite (Gorton and Johansson, 1980), while the potential of the
protein-bound FAD is expected to be different. This also supports the
assumption that the FAD signal in Fig. 1A rather stems from FAD
that has been released from the active site than the enzymebound FAD.
The FAD redox peaks scale linearly with the potential scan rate
(Fig. 1B) thus confirming a surface-confined redox process (Bard
and Faulkner, 1980) characterized by the ET rate constant ks of
83 s 1 as calculated through the Laviron formalism (Laviron,
1979) (Fig. 1C). The peak potentials in CV and DPV linearly
depended on pH with a slope of 60 77 mV per pH unit
(Fig. S2, ESI), indicating an equal number of protons and electrons
involved in the redox process (Bard and Faulkner, 1980) in
consistence with other reports on the electrochemistry of FAD/
FADH2 (Gorton and Johansson, 1980; Wei and Omanovic, 2008).
The analysis of the full peak width at the half-peak height for a
221
cathodic and anodic process (Laviron, 1979) at low scan rates
confirmed a 2e transformation of the FAD/FADH2 couple with
the transfer coefficient a of 0.42.
In the presence of oxygen, its electrochemical reduction
started from approximately 170 mV (Fig. 2, inset), correlating
with electrochemical reduction of O2 catalyzed by the heme of
HMP (on bare Gr electroreduction of O2 starts at potentials lower
than 300 mV). This electrocatalytic reduction reaction is characteristic of the heme and heme proteins such as peroxidases and
different type of globins (Ferapontova, 2004); it may be also
observed with more complex heme-containing enzymes such as
theophylline oxidase (Shipovskov and Ferapontova, 2008), cytochromes P450 (Uldt et al., 2004) and other types flavohemoglobins, e.g. from Ralstonia eutropha entrapped into the
methylcellulose film (solely the heme redox transformation and
no FAD signals were detected) (de Oliveira et al., 2007).
Upon addition of NADH no catalytic oxidation of the substrate
was observed at potentials of the FAD/FADH2 redox couple, but
only at potentials higher than 50 mV, which correlated well with
catalysis by the surface quinone groups (Fig. 3). This suggests the
ability of surface quinones (also in the absence of HMP) to
mediate the NADH oxidation, alas with very slow kinetics: the
oxidation current was practically independent on the NADH
concentration within the quinone peak potential window. At
more positive potentials, non-catalytic electrochemical oxidation
of NADH proceeded with a higher rate. The lack of catalysis at the
potentials corresponding to the observed FAD/FADH2 redox peaks
is believed to be caused by two factors. Firstly, the orientation of
HMP at the electrode is inappropriate for catalysis—HMP is
orientated predominantly through the FAD domain (where the
catalytic conversion of a substrate should occur), rather than
through the heme domain. Secondly, denaturation of the protein
occurs at the electrode surface, which is accompanied by the
aforementioned partial loss of the FAD cofactor.
3.2. Bioelectrocatalysis of HMP wired by the redox polymers
To achieve bioelectrocatalysis of NADH oxidation by HMP,
the protein was co-immobilized with a redox mediator: the
active sites of the protein were ‘‘wired’’ to the electrode surface
through the Os-complexes bound to the polymer matrix of two
types (Fig. 1S, ESI). The used Os complexes contained either two
50
[NADH] , mM
0
0.62
1.25
1.9
2.5
40
30
20
10
0
-10
-20
-30
-40
-0.8
-0.6
-0.4
-0.2
0.0
0.2
0.4
E vs Ag/AgCl , V
Fig. 3. Representative CVs recorded with the HMP-modified graphite electrode in
deoxygenated 20 mM Tris buffer, pH 7.4, in the presence of varying concentration
of NADH (as indicated); scan rate 20 mV s 1.
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M. Sosna et al. / Biosensors and Bioelectronics 42 (2013) 219–224
Fig. 4. Representative cyclic voltammograms of HMP ‘‘wired’’ to the electrode by redox polymer I (left panel) and redox polymer II (right panel) in the presence of varying
concentrations of NADH (concentrations, in mM, indicated in the graphs); scan rate 30 mV s 1. Insets: the dependence of catalytic currents of NADH oxidation on the
concentration of NADH.
4,40 -dimethyl-2,20 -bipyridine ligands or two N,N0 -methyl-2,20 -biimidazole ligands, for polymers I and II, respectively. The remaining Os coordination sites were occupied by the nitrogen atom of
imidazole bound to the polymer backbone and chloride ion
(polymer I), and 2,20 -biimidazole moiety linked to the polymer
backbone (polymer II). Both types of Os complexes are cationic
and therefore can bind electrostatically to the anionic domains of
HMP (pI r5.0), which are predominantly located in the vicinity of
the NADH-binding domain and on the molecular surface of the
heme containing domain (Ilari et al., 2002). Either type of the
polymeric units was mixed with HMP in the presence of the
bifunctional crosslinking agent PEGDGE to form a three dimensional
redox-polymer matrix with the redox protein incorporated into it,
which therewith remained immobilized at the electrode surface.
In the absence of NADH, CVs of HMP entrapped into the
polymer I matrix showed two distinct sets of redox peaks
(Fig. 4, left panel). Similar to the HMP-modified electrodes, a set
of peaks corresponding to the FAD/FADH2 redox transformation
was seen at approximately 470 mV. At more positive potentials
(18973 mV) another redox couple matching the oxidation/
reduction potentials of the polymer bound Os centers was
observed. Upon addition of NADH the flavin oxidation peak
remained unchanged thus indicating that HMP is not capable of
unmediated bioelectrocatalysis when it is entrapped into the
polymeric matrix, similarly to the results obtained with HMP just
adsorbed on graphite. In the presence of NADH, its catalytic
oxidation can be followed at potentials corresponding to the Os
complex redox transformation. The CVs exhibited a large
enhancement in the anodic peak current with a concomitant
decrease in the cathodic peak current, indicating a strong catalytic
activity of HMP in the reaction of NADH oxidation, mediated by
Os centers wiring the electron transfer between the electrode and
the protein (Fig. 4, left panel). Similarly to previously reported
results (Antiochia and Gorton, 2007; Tasca et al., 2008), no
electrocatalytic oxidation of NADH by Os polymer in the absence
of the biocatalyst was observed, namely when HMP was not
entrapped into the polymer matrix.
Thus, in the presence of the Os polymer mediator, replacing O2
as a natural electron acceptor, HMP demonstrated its ability to
electrocatalytically oxidize NADH. Redox polymer I, mediating
the ET reaction between the electrode and protein, allowed a
noticeable reduction of the NADH oxidation overpotential when
compared to the unmodified carbon electrodes. However, to avoid
possible interfering oxidations (Fukuda et al., 2008) a further
decrease in the overpotential was achieved by using redox
polymer II, which contains Os centers with a lower redox
potential, as mediator.
As can be seen in Fig. 4 (right panel), a couple of redox peaks
from the Os complex centered at 20678 mV can be seen in CVs
recorded with the HMP-polymer II-modified electrode in NADHfree solutions, which is in accordance with previous reports using
this polymer (Mao et al., 2003; Timur et al., 2006). Upon addition
of NADH, bioelectrocatalytic oxidation of NADH occurred at a
significantly reduced overpotential, with onset corresponding to
the redox potential of the Os complex, which mediates the ET
reaction between the electrode and the protein. At a given
concentration of NADH the catalytic currents were noticeably
lower when compared to the NADH oxidation at electrodes
modified with HMP and polymer I. Consistently with earlier
reports (Fukuda et al., 2008), the higher redox potential of
polymer I results in higher rate of NADH oxidation: the rate
constant of the reaction between HMP and the mediator increases
exponentially with increasing difference in their redox potentials
(based on the ET rate constant–free energy relationship (Bard and
Faulkner, 1980)).
Fig. 4 shows NADH-oxidation calibration plots constructed for
both polymers in order to estimate the apparent efficiency of
bioelectrocatalytic oxidation of NADH. The apparent Michaelis
constants, Kapp
were found to be 10.8 and 9.5 mM for polymer
M
I- and polymer II-modified HMP electrodes, respectively, suggesting that in general, interactions of NADH with enzyme bound FAD
and the stability of the intermediate complex remain almost the
same, i.e. the overall fold of HMP and in particular the local
chemical environment around the FAD binding domain is unaffected by the polymer matrix.
The values of Imax, correlating with the overall rate of oxidation
at excess of NADH, differ appreciably (38 and 23 mA for polymers I
and II, respectively) and point to the oxidation of the reduced
form of HMP by Os mediator as a possible rate determining step.
Redox polymer I has significantly higher redox potential and
therefore the driving force for oxidation should result in a higher
rate of oxidation of the FADH2 cofactor. However, the overall
M. Sosna et al. / Biosensors and Bioelectronics 42 (2013) 219–224
oxidation currents also depend on the concentration of Os centers
(9.570.5 nmol cm 2 for polymer I and 1.5 70.3 nmol cm 2 for
polymer II, as determined from the low potential scan rate CVs)
and their diffusivity within the film (Bartlett and Pratt, 1995; Mao
et al., 2003), provided sufficient flexibility of the polymer tethers.
Finally, the overall rate depends also on the amount of the intact
HMP in the film, which is not known, since the non-turnover
flavin voltammetric response can originate both from the intact
and partially de-natured (and thus catalytically inactive) protein.
Due to the complexity of the system the currently available
data do not allow the precise identification of the rate determining step or its kinetics. The most important is the demonstrated
ability of HMP to bioelectrocatalytically oxidize NADH at reduced
overpotentials in the presence of the appropriate ET mediators.
The HMP modified electrodes may be favorably compared to
other enzyme based electrodes for NADH oxidation. A jmax of
325 mA cm 2 (at 0.1 V) for the polymer II-HMP system significantly exceeds those obtained by (Kobayashi et al. (1992)) using
diaphorase (1 mA cm 2 at 0 V) and ferredoxin NADP þ reductase
(0.9 mA cm 2 at 0 V) and by (Barker et al. (2007)) with subcomplex of Complex I (NADH:ubiquinone oxidoreductase)
(90.1 mA cm 2 at 0.2 V). It has to be noted, however, that in
both of these works enzyme electrodes based on direct ET were
used and the reference potentials differed, thus making a direct
comparison not straightforward. Among comparable systems,
NADH oxidation by diaphorase wired by the low-potential Os
polymer to graphite electrodes with its jmax of 57 mA cm 2 (at
0.1 V) (Tasca et al., 2008) allows us to consider HMP as a
promising future component of bioanalytical and bioenergetic
systems involving NADH oxidation/recycling.
4. Conclusions
In the present work E. coli flavohemoglobin HMP was electrochemically characterized and its potential as a NADH oxidation
biocatalyst in electrochemical biosensors and biofuel cell components was studied. HMP adsorbed on graphite displayed redox
signals both from its FAD and heme moieties however, no
bioelectrocatalysis of NADH oxidation was observed at potentials
of the individual enzyme redox components. This might result both
from the partial denaturation of HMP accompanied by the release
of the FAD cofactor and/or orientation of HMP unfavorable for
bioelectrocatalysis. HMP incorporated in Os-complex redox polymer matrices and by this means wired to the electrodes through
the Os-redox relay was able to bioelectrocatalytically oxidize
NADH at reduced overpotentials corresponding to the redox
transformation of the specific Os complex. NADH oxidation by
HMP incorporated within the low potential polymer II occurred at
a potential only 300 mV more positive than the formal potential
of the NAD/NADH redox couple (0.52 V vs. Ag9AgCl) (Voet and
Voet, 2004) and the potential of FAD in HMP. It is currently unclear
whether the Os centers oxidize FADH2 directly or with involvement of the heme oxidation followed by intramolecular ET. The
latter seems to be more likely as the proximity of Os complex to
the FAD domain would cause a steric hindrance for the NADH
substrate molecules. It is also consistent with the HMP behavior in
homogeneous conditions. Further work is focused on more detailed
studies of the presented bioelectrocatalytic system.
Acknowledgments
The work was supported by the Danish Council for Independent
Research, Natural Sciences (FNU), Project number 11-107176, and
The Swedish Research Council, Project number 2010-5031.
223
Appendix A. Supporting information
Supplementary data associated with this article can be found in
the online version at http://dx.doi.org/10.1016/j.bios.2012.10.094.
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