NAD-dependent dehydrogenase bioelectrocatalysis: The ability of a naphthoquinone redox polymer to regenerate NAD

Article abstract of DOI:10.1039/c5cc09161f

Electron mediation between NAD-dependent enzymes using quinone moieties typically requires the use of a diaphorase as an intermediary enzyme. The ability for a naphthoquinone redox polymer to independently oxidize enzymatically-generated NADH is demonstrated for application to glucose/O₂ enzymatic fuel cells.

Full text of DOI:10.1039/c5cc09161f

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NAD-dependent dehydrogenase bioelectrocatalysis:
the ability of a naphthoquinone redox polymer to
regenerate NAD†
Cite this:DOI: 10.1039/c5cc09161f
Received 4th November 2015,
Accepted 24th November 2015
Sofiene Abdellaoui, Ross D. Milton, Timothy Quah and Shelley D. Minteer*
DOI: 10.1039/c5cc09161f
www.rsc.org/chemcomm
Electron mediation between NAD-dependent enzymes using quinone
While phenothiazines (such as methylene blue, methylene
moieties typically requires the use of a diaphorase as an intermediary green, thionine and toluidine blue) are capable of oxidizing
2
2–24
enzyme. The ability for a naphthoquinone redox polymer to indepen- NADH without the need for diaphorase,
their immobilization
dently oxidize enzymatically-generated NADH is demonstrated for by electropolymerization or covalent attachment to polymers
application to glucose/O
2
enzymatic fuel cells.
results in a positive potential shift to less-desired onset potentials
of NADH oxidation, due to the inherent change in chemical
Enzymatic fuel cells (EFCs) are devices that utilize enzymes at their structure of the dye. Additionally, the purification of resulting
anodes (and commonly at their cathodes) to facilitate the produc- phenothiazine polymers is not trivial whereby many polymers
tion of electrical energy from common energy-dense fuels, such as still possess entrapped non-covalently modified phenothiazine.
1–4
sugars and alcohols. To date, high performance EFCs frequently
We recently reported a naphthoquinone redox polymer
incorporate flavin adenine dinucleotide-dependent (FAD) enzymes (NQ-LPEI) that was able to mediate electrons between FAD-GDH
(
1
i.e. FAD-dependent glucose dehydrogenase (FAD-GDH, E.C.: and yield an EFC with a remarkably high maximum current density
À2
À2
.1.99.10) and glucose oxidase (GOx, E.C.: 1.1.3.4)) commonly (5.4 mA cm ) and maximum power density (2.3 mW cm at
6
coupled to a single electron mediator; however, a large portion 0.55 V). Herein, we report the ability of the NQ-LPEI redox
of EFCs make use of nicotinamide adenine dinucleotide- polymer to directly oxidize enzymatically-generated NADH
dependent (NAD) enzymes (i.e. NAD-dependent glucose dehy- without the use of diaphorase. This NQ-LPEI redox polymer also
5
–9
drogenase (NAD-GDH, E.C.: 1.1.1.47)).
offers additional benefits of low cost (compared to PQQ-based
electron media- systems) and the ability to immobilize NAD-dependent enzymes
1
0–12
13–16
Alongside ferrocene
and osmium
tors, quinones (typically naphthoquinones and anthraqui- within a 3-dimensional network. A glucose/O EFC was constructed
2
nones) have been coupled with FAD- and NAD-dependent using NAD-GDH at the bioanode and laccase at the biocathode,
enzymes as efficient electron mediators, however an additional yielding maximum current and power densities of 505.7 Æ
À2
À2
enzyme (diaphorase) is required as an intermediary between 56.7 mA cm and 123.2 Æ 16.3 mW cm (respectively), with
5
,6,17
NAD-dependent enzymes and quinone electron mediators.
ortho-Quinones have been reported to oxidize NADH,
an associated OCP of 766.7 Æ 4.9 mV.
Scheme 1 outlines the mechanism by which bioelectrocata-
ranging from simple catechol derivatives to the enzymatic lytic glucose oxidation by NAD-GDH is ultimately facilitated by
1
8,19
cofactor pyrroloquinoline quinone (PQQ).
Further, Katz the NQ redox polymer to the electrode surface. As previously
et al. demonstrated the ability of an immobilized PQQ reported, the NQ-LPEI redox polymer makes use of self-
À11
2
À1
monolayer to efficiently oxidize NADH at gold and platinum exchange (electron diffusion coefficient of 1.3 Â 10
cm s )
2
0
electrode surfaces. Another example is a ruthenium electron to permit improved electron transport whereby the oxidation of
mediator recently published by Cosnier et al. that possessed a NADH is not limited to a single adsorbed enzyme/mediator layer
9
,10-phenanthraquinone ligand and was able to directly oxidize (such as that typically offered by p–p stacking architectures) but
2
1
6
enzymatically-generated NADH.
rather an expanded 3D electrode architecture.
Fig. 1A presents cyclic voltammetry demonstrating the ability
of the NQ-LPEI redox polymer to oxidize NADH, whereby NADH
oxidation was initially investigated by the addition of reduced
NADH to the citrate/phosphate buffer. It is important to demon-
strate that the oxidation of NADH yields a biologically compatible
Departments of Chemistry and Materials Science and Engineering,
University of Utah, 315 S 1400 E, Room 2020, Salt Lake City, UT, 84112, USA.
E-mail: minteer@chem.utah.edu
†
Electronic supplementary information (ESI) available: Experimental procedures,
+
Fig. S1 (Michaelis–Menten kinetics). See DOI: 10.1039/c5cc09161f
NAD cofactor, thus, the ability of the bioelectrode to oxidize
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Fig. 2 Overlaid pH profile of electrocatalytic NADH oxidation (5 mM)
at 0 V (vs. SCE) at NQ-LPEI/NAD-GDH bioelectrodes and NAD-GDH
+
activities monitored by UV-Vis at 340 nm in presence of 5 mM NAD
Scheme 1 Bioelectrocatalytic oxidation of glucose by NAD-dependent
glucose dehydrogenase.
and 100 mM glucose.
NADH generated from glucose oxidation by NAD-GDH was optimal pH of glucose oxidation by NAD-GDH (as determined
investigated (Fig. 1B). On Toray paper electrodes, a clear by UV-Vis experiments) and near-neutral NADH oxidation by
oxidative wave following the addition of glucose was observed. the NQ-LPEI polymer.
Control bioelectrodes comprised of an alternative LPEI-based
Amperometric experiments were employed to determine the
polymer (C -LPEI, where the NQ moiety is replaced by a optimal concentration of NAD /NADH to be used with the
+
8
8
-carbon chain) and denatured NAD-GDH confirmed that the NQ-LPEI/NAD-GDH bioelectrodes (Fig. 3A) at 0 V (vs. SCE at
activity observed is in fact due to the enzymatic oxidation pH 6.5). The Michaelis–Menten model (by nonlinear regres-
of glucose coupled to the enzymatic reduction of NADH (data sion) was employed in an attempt to evaluate the apparent
not shown).
Enzymatic activity assays and cyclic voltammetry were polymer and a control polymer (C
employed to determine the optimal pH for glucose oxidation and Toray paper electrodes. An apparent Michaelis constant
kinetic parameters of NADH oxidation by the NQ-LPEI redox
8
-LPEI), on both glassy carbon
app
À2
by NAD-GDH in a homogeneous setting and the optimal pH (K
M
) of 12.0 Æ 0.6 mM NADH and a Jmax of 568.3 Æ 14.6 mA cm
for NADH oxidation by the NQ-LPEI redox polymer, respec- were obtained for NQ-LPEI/NAD-GDH bioelectrodes on glassy
tively (Fig. 2). Following electrochemical characterization was carbon electrodes. The K
performed at pH 6.5, as a compromise between the alkaline when the bioelectrodes were scaled up to Toray paper, with an
app
M
decreased to 4.3 Æ 0.2 mM NADH
Fig. 1 (A) Representative cyclic voltammograms of NQ-LPEI/NAD-GDH bioelectrodes in citrate/phosphate buffer (0.2 M, pH 6.5) in the absence (black
2
dashed line) and presence (black solid line) of 50 mM NADH, prepared on Toray paper electrodes (0.25 cm ). Control bioelectrodes were prepared
8
with C -LPEI/NAD-GDH (gray dashed line) in the presence of 50 mM NADH, on Toray paper electrodes. (B) Representative cyclic voltammograms of
+
NQ-LPEI/NAD-GDH bioelectrodes in citrate/phosphate buffer (0.2 M, pH 6.5) containing 10 mM NAD , in the absence (black dashed line) and presence
of 200 mM glucose (black solid line). Black lines represent bioelectrodes prepared on Toray paper, whereas the gray line represents electrodes prepared
À1
on glassy carbon electrodes. All electrodes were evaluated in hydrostatic quiescent buffer at a scan rate of 5 mV s (n = 3, error bars represent one
standard deviation).
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Fig. 3 (A) Calibration curve for the oxidation of NADH by the NQ-LPEI/NAD-GDH bioelectrodes at 0 V (vs. SCE). Bioelectrodes were prepared on glassy
carbon (black) and Toray paper electrodes (red). Control bioelectrodes prepared with C -LPEI were evaluated on glassy carbon (gray) and Toray paper
8
electrodes (white). (B) Apparent Michaelis–Menten kinetics for the bioelectrocatalytic oxidation of glucose by the NQ-LPEI/NAD-GDH bioelectrodes
+
at 0 V (vs. SCE) in the presence of 10 mM NAD . Bioelectrodes were prepared on glassy carbon (black) and Toray paper electrodes (red). Control
bioelectrodes prepared with C
8
-LPEI were evaluated on glassy carbon (gray) and Toray paper electrodes (white). All electrodes were evaluated at in
stirred citrate/phosphate buffer (0.2 M, pH 6.5) with no additional gas removal/purging (n = 3, error bars represent one standard deviation).
À2
increased J
of 756.9 Æ 12.0 mA cm . The decreased bioelectrodes for glucose oxidation at 0 V (vs. SCE at pH 6.5,
max
app
K
reflects an increase in affinity for NADH oxidation by Fig. 3B). Apparent Michaelis–Menten kinetics were determined
M
app
M
the Toray paper bioelectrodes, also resulting in an increased by non-linear regression and a K
of 3.0 Æ 0.1 mM glucose
J
max; thus, further electrochemical characterization was per- was determined, with an associated Jmax of 238.5 Æ 1.9 mA cmÀ2
,
formed on Toray paper electrodes. Small contributions from for NQ-LPEI/NAD-GDH bioelectrodes prepared on Toray paper
the ability of the MWCNTs to electro-oxidize NADH (using the electrodes. Interestingly, bioelectrodes prepared on glassy carbon
À2
app
M
C
8
-LPEI control polymer) result in a Jmax of 154.0 Æ 1.4 mA cm
,
electrodes had a K
of 2.7 Æ 0.1 mM glucose, with an associated
À2
which is approximately 20% of the catalytic current obtained Jmax of 184.3 Æ 1.5 mA cm . Determination of the apparent
for the NQ-LPEI redox polymer.
Michaelis–Menten kinetics by Lineweaver–Burk double recip-
+
app
Since high concentrations of NAD /NADH are relatively rocal linearization yielded a KM of 3.1 Æ 0.1 mM glucose, with
+
À2
costly, a concentration of 10 mM NAD was selected for further an associated Jmax of 244.0 Æ 16.2 mA cm
.
evaluation. Control experiments were performed using a compa-
UV-Vis experiments determined a comparable solution-based
app
rable LPEI-based polymer (absent of the NQ moiety, C
both glassy carbon and Toray paper electrodes.
8
-LPEI), on
K
M
of 3.0 Æ 0.2 mM glucose and a Vmax of 10.1 Æ 0.1 mU, by
following the production of NADH at 340 nm (Fig. S1, ESI†).
Amperometric experiments were also used to evaluate the Additional UV-Vis experiments were performed to confirm
apparent enzymatic kinetics of the final NQ-LPEI/NAD-GDH the ability of NQ to oxidize NADH, where the reduction of a
2
Fig. 4 Polarization and corresponding power curve for a glucose/O EFC constructed using a NQ-LPEI/NAD-GDH bioanode coupled with a laccase
DET-type biocathode. The EFC was operated using a proton exchange membrane to separate the anodic and cathodic chambers, to prevent the
crossover and resulting oxidation of NADH at the biocathode. Thus, the anodic chamber consisted of citrate/phosphate buffer (0.2 M, pH 6.5) containing
2
00 mM glucose, whereas the cathodic chamber did not contain glucose (citrate/phosphate buffer, 0.2 M, pH 5.5). The anodic and cathodic chambers
À1
were stirred. Polarization was performed by galvanostatically drawing increasing current from the EFC at a slow sweep rate (0.1 mA s ), until short circuit.
Current-limiting bioanodes were prepared on Toray paper electrodes with a geometric surface area of 0.25 cm , whereas biocathodes were prepared on
Toray paper electrodes with a geometric surface area of 1 cm .
2
2
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water-soluble naphthoquinone (1,2-naphthoquinone-4-sulfonic Individual Fellowship (Global) under the EU Commission’s
app
M
acid sodium salt) yielded a K
of 2.9 Æ 0.1 mM glucose and Horizon 2020 framework (project number 654836).
a Vmax of 10.6 Æ 0.1 mU (followed at 450 nm) (Fig. S1, ESI†).
The similarities between the kinetics of the immobilized
NAD-GDH and the solution based NAD-GDH reflect the limitation
of the NAD-GDH on the bioelectrode by the concentration of
NADH (as outlined above), resulting in a system that can be
used to further study the enzymatic kinetics of NAD-dependent
enzymes.
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Fig. 4 presents a polarization curve and corresponding power
curve for a representative EFC; experiments were performed in
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associated maximum current and power densities of 505.7 Æ
À2
À2
5
6.7 mA cm and 123.2 Æ 16.3 mW cm , respectively. Blank
EFCs (tested in the absence of glucose) yielded approximate
power densities of 17 mW cm , produced as a function of the
À2
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of diaphorase, resulting in a simplified system. Resulting
NAD-GDH bioelectrodes were utilized as glucose bioanodes
with laccase O -reducing biocathodes in a glucose/O EFC.
It is anticipated that the system presented within this manuscript
will impact a large number of NAD-dependent EFCs and cascade
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The authors thank the National Science Foundation and the
Army Research Office MURI (#W911NF1410263) grant for funding.
RDM acknowledges funding from a Marie Curie-Skłodowska
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