DSpace at VNU: Flexible, micron-scaled superoxide sensor for in vivo applications

Journal of Electroanalytical Chemistry 662 (2011) 100–104

Contents lists available at ScienceDirect

Journal of Electroanalytical Chemistry
journal homepage: www.elsevier.com/locate/jelechem

Flexible, micron-scaled superoxide sensor for in vivo applications
Rebekah C.K. Wilson a, Dao Thanh Phuong b, Edward Chainani a, Alexander Scheeline a,⇑
a
b

Department of Chemistry, University of Illinois at Urbana-Champaign, 600 South Mathews Avenue, Urbana, IL 61801, United States
Faculty of Chemistry, Hanoi University of Science, VNU 334 Nguyen Trai, Thanh Xuan, Ha Noi, Viet Nam

a r t i c l e

i n f o

Article history:
Available online 29 March 2011
Keywords:
Superoxide sensor
Flexible microelectrode
Chronoamperometry
Reactive oxygen species sensor
Cytochrome c immobilized protein electrode
Thiol on gold modified electrode

a b s t r a c t
Superoxide radical plays an important role in cell signaling. However, certain events can result in a large

increase in superoxide concentration which has been linked to, among other conditions, inflammation,
neurodegenerative diseases, and cancer. Consequently, in vivo detection of superoxide is of great interest.
Previously, due to brittleness, instability, or size, superoxide sensors have been limited in their ability for
in vivo work. We report the development of a flexible, micron-scale, superoxide sensor. Thin gold films
are patterned on Kapton™ to form multiple electrodes that constitute the sensor. Cytochrome c was covalently anchored to the working electrode using a self-assembled monolayer of 3,30 -Dithiodipropionic acid
di(N-hydroxysuccinimide ester). Calibration showed a linear response within the constraints imposed by
using xanthine/xanthine oxidase as the superoxide source. Testing demonstrated that interference from
physiological levels of NADH, citric acid, and uric acid to be insignificant. However, minor interference
was seen in the presence of H2O2 and glucose, and significant interference arose from ascorbic acid, a
known radical scavenger. Qualitative observations provide insight into the preparation and cleaning of
thin layer gold on Kapton™.
Ó 2011 Elsevier B.V. All rights reserved.

1. Introduction
Reactive oxygen species (ROS), or pro-oxidants, are molecules
or ions formed by the reduction of oxygen and are highly reactive.
These species include, but are not limited to, singlet oxygen (1O2),
0
superoxide (OÅÀ
2 ), peroxides (R-O-O-R ), the hydroxyl radical (OH),
and hypochlorous acid (HOCl). They can be generated via photochemistry and by toxic chemical or drug exposure. Biologically,
OÅÀ
2 is the result of a one-electron reduction of molecular oxygen
during cell metabolism and also plays an important role in cell signaling [1,2]. Enzymes, such as superoxide dismutase (SOD), and
antioxidants, such as ascorbic acid, are present biologically to keep
the concentration of superoxide manageable. However, during
times of environmental stress, the levels of OÅÀ
2 can exceed those
which can be removed by natural defenses and can lead to destruction of vital cell structures, leading ultimately to cancer, ischemia/
reperfusion damage, diabetes, aging-correlated pathology, or cardiovascular disease [3–5].

Because of its damaging effects, the detection and quantification of OÅÀ
2 has been the subject of biomedical interest for decades
[6–11]. A robust and reliable detection scheme for superoxide
would lead to better understanding of its effects. However, the
detection of OÅÀ
2 is complicated by its rapid dismutation. At physi-

⇑ Corresponding author. Tel.: +1 217 333 2999; fax: +1 217 265 6290.
E-mail address: (A. Scheeline).
1572-6657/$ - see front matter Ó 2011 Elsevier B.V. All rights reserved.
doi:10.1016/j.jelechem.2011.03.024

ological pHs, the half-life ranges from milliseconds to seconds [12].
Therefore detection of OÅÀ
2 also requires sensitivity and selectivity
along with rapid detection for in vitro and in vivo applications. Previous studies have employed numerous analytical techniques to
explore OÅÀ
reactions. Among these, electron spin resonance
2
(ESR) has been a reliable way to monitor ROS, but requires long
averaging times, and is restricted in sensitivity [13]. High performance liquid chromatography (HPLC) is another popular form of
ROS detection, but requires digestion of the biological samples,
which is not compatible with in vivo work [14,15]. Chemiluminescence also offers specific targeting of certain ROS [16–18] However,
the chemicals involved may easily disrupt the biological system
one is attempting to monitor, or may interfere with the reaction
to oxidative stress.
Electrochemical detection of OÅÀ
2 is promising not only for its
biocompatibility, fast response times, and selectivity, but can also
be easily integrated into portable devices. Exploiting the biological

function of proteins by integrating them onto, e.g., a gold surface
allows for selectivity. Cytochrome c (cyt c) is a heme protein that
can be found within the inner membrane of mitochondria. It can
function as a catalyst for hydroxylation and aromatic oxidation,
and is an initiator in apoptosis. Cytochrome c also can be reduced
by OÅÀ
2 and can be anchored to the surface of an electrode using a
promoter species. Until now, OÅÀ
2 electrodes have been engineered
in such a way that the working area is much too large or the support too brittle for some in vivo work, limiting its capabilities. Further, calibration of such a sensor has not been found in literature


R.C.K. Wilson et al. / Journal of Electroanalytical Chemistry 662 (2011) 100–104

[19–24]. Presented here is the first micron-scale flexible OÅÀ
2 sensor
compatible for in vivo detection. Here, patterned gold electrodes on
a Kapton™ surface were modified using cyt c anchored by a selfassembled monolayer of dithiobis-(succinimidyl) propionate
(DTSP). Using the xanthine/xanthine oxidase system to produce
OÅÀ
2 , the signal response of this sensor is presented along with calibration and exposure to biological interferents.
Electrodes were fabricated such that working, counter, and reference electrodes would all fit on a 200 lm wide projection a few
millimeters long, as initial applications envisioned included study
of noise-induced hearing loss [25–27] in small mammals and monitoring of reactions in ultrasonically-levitated drops [28–30].

2. Materials and methods
2.1. Materials
Unless noted, all chemicals are of reagent grade, used as received. Mono- and dibasic potassium phosphate, oxalic acid,
hydrogen peroxide, potassium carbonate, 3,30 -Dithiodipropionic
acid di(N-hydroxysuccinimide ester) (DTSP), xanthine, xanthine

oxidase, superoxide dismutase (SOD), b-nicotinamide adenine
dinucleotide, reduced disodium salt (NADH), citric acid, ascorbic
acid, glucose, cyt c and organic solvents were obtained from Sigma
Aldrich (St. Louis, Mo) Kapton™ 500 FCP and 500 VN, both 127
micrometers thick (DuPont) were used for the sensor substrate.
Lithography was achieved using S 1805 positive photoresist
(Microchem), MF 319 developer (Shipley), and GE 8111 gold etch
(Transene).
A 50 mM stock, pH 7.4 phosphate solution was created by dissolving 0.1558 g of monosodium phosphate, monohydrate and
1.037 g disodium phosphate, heptahydrate into 100 mL of deionized (DI) water. The stock solution was further diluted to 10:1 to
obtain a 25 mM solution, which was used for experimentation.
A 2 mM xanthine solution was made by dissolving 0.006 g of
xanthine in 50 mL of 25 mM phosphate buffer (pH 7.4) and was
mechanically stirred on low at low heat to fully dissolve the xanthine. The solution is allowed to cool to room temperature before
being used. Over time, the xanthine precipitates and can be redissolved by heat and stirring.
A 30 mM solution of SOD was made by dissolving 0.2 mg of SOD
in 200 mL of 25 mM phosphate buffer and kept on ice.
A 0.8 mg/mL solution of xanthine oxidase was made by diluting
5 mL of a 16 mg/mL suspension of xanthine oxidase to 105 lL
using 25 mM phosphate buffer (pH 7.4) and kept on ice.
Artificial perilymph (a mixture chosen for its relevance to future
studies of noise-induced hearing loss) contained 112 mM NaCl,
4.2 mM KCl, 1 mM MgCl2, 3.4 mM NaH2PO4, 5.3 mM Na2HPO4,
21 mM NaHCO3 M and 3.6 mM glucose in DI water. Specifically,
0.659 g NaCl, 0.0313 g KCl, 0.0203 g MgCl2 Á 6H2O, 0.0475 g
NaH2PO4 Á H2O, 0.0764 g Na2HPO4, 0.1764 g NaHCO3, and
0.0649 g glucose for every 100 mL of DI water.
Unless otherwise noted, a CH660 (CH Instruments, Austin, TX)
was the potentiostat used. When indicated, a Keithley 6485
Picoammeter (Solon, OH) was used. All chronoamperometry data

were collected at 3 Hz using the digitizer built into the picoammeter. Unless stated, all electrochemical measurements were made
using a Ag/AgCl reference (CH Instruments, Austin, TX, [ClÀ] = 3 M)
electrode and all potentials are reported as such. A Pt wire was
used as a counter electrode. All ebeam work is done on a Temescal
six pocket E-Beam Evaporation System (Livermore, CA). All UV
exposures were completed using a Karl Suss MJB3 Mask aligner
(SUSS MicroTec Inc., Waterbury, VT). A HI3222 pH/ORP/ISE meter
(Hanna Instruments, Woonsocket, RI) was used for all pH measurements. Oxygen plasma cleaning is done with a MARCH CS 1701 RIE

101

and RFX 600 generator (Nordson MARCH, Carlsbad,CA). All nitrogen was from tank sources, not house supply.
The platform of this sensor is inexpensive and disposable with
the intention to calibrate, use, validate and dispose without reuse.
Therefore, all reported experiments use each electrode for only one
experiment.
2.2. Fabrication and modification of sensor
Kapton™ surfaces were cleaned prior to metal deposition by
successive washing in acetone, ethanol and isopropyl alcohol, followed by drying with N2. The substrates were then exposed to oxygen plasma (80 sccm, 500 mTorr, 200 W) for 300 s and placed
immediately into the ebeam deposition chamber. A 60–80 Å adhesion layer of Ti was coated (1 Å sÀ1) followed by 2000 Å of Au. Upon
removal from the deposition chamber, the newly coated surfaces
were flushed with N2 and placed in a dessicator. Typical lithography procedures were used to pattern gold electrodes [31]. Specifically, S1805 positive photoresist was spun onto the surface of the
gold (300 rpm, 30 s) and baked at 110 °C for 3 min. Once removed,
it was allowed to cool. A positive mask was placed onto the surface
and held in place with a quartz slide. The surface was exposed to
UV light (12 s, 300 W), placed in MF 319 developer (5–10 s) and
quickly rinsed with DI water. Ti and Au were etched in GE 8111
(2–4 min) and rinsed generously with DI water. Photoresist was removed using acetone. Due to the precise dimensions required, laser cutting was used to excise the sensor from the Kapton™
sheet. Sensors were kept in a desiccator until modification to reduce delamination of Au.
Areas where isolation from the environment is necessary, the

gold surfaces between sensing and contact pad areas, were coated
by hand-painting polyimide onto the surface and cured at 100 °C
for 1 h. The working area was 3 mm by 50 lm but could have been
made smaller by isolating more of the gold lead. The electrode surface is then cleaned by placing it in oxygen plasma for 2–4 min before being chemically modified. Typical electrochemical cleaning in
acidic solutions causes the gold to delaminate from the Ti or Cr
adhesion layer. Covalent binding of cyt c was largely based on a
procedure from Chen et al. [32]. Briefly, a clean gold electrode
was placed in a 10 mM (dried over sieves DMSO) solution of DTSP
for 2 h at room temperature. Once removed, the electrode was
lightly rinsed with dry DMSO, then DI water and dried with nitrogen. Gold surfaces where DMSO is not desired can be placed in
0.5 M KOH (methanol) and exposed to À1.5 V for 120 s and immediately rinsed with DI water. The electrode was then placed in a
2 mg/mL solution of cyt c (25 mM phosphate buffer, pH 7.4) at
4 °C for 24 h. DTSP contains the NHS ester which provides a good
leaving group allowing amines on the cyt c to covalently bind to
the DTSP-modified surface. Once removed from the cyt c solution,
it was lightly rinsed with cold phosphate buffer and stored in buffer at 4 °C when not in use.
2.3. Calibration of superoxide sensor
The DTSP/cyt c modified working electrode was removed from
25 mM phosphate buffer (pH 7.4), rinsed with cold 25 mM phosphate buffer (pH 7.4), and placed in a 2 mL solution of xanthine
(2 mM). A stir bar was added to the reaction cell along with a
Ag/AgCl reference electrode and a Pt wire as the counter electrode.
A potential was applied to the counter electrode using a potentiostat so that the working electrode was at 200 mV (to ensure proper
oxidation of cyt c) with respect to the reference electrode. The
working electrode was connected to the picoammeter to measure
current. Background current was recorded for a period of time to
establish a stable background level. Serial injections of the xanthine oxidase solution (20 lL) were added to the reaction cell


102

R.C.K. Wilson et al. / Journal of Electroanalytical Chemistry 662 (2011) 100–104

4. Results and discussion

followed by sonication in H2SO4. Alumina powder is too abrasive
for thin films. Piranha (1:3, H2O2:H2SO4) creates too harsh of an
environment, causing Kapton™ to warp and gold to delaminate.
A survey of literature using gold patterned electrodes on Kapton™
found either vague or no instructions for cleaning the gold surfaces
prior to being modified [33–35]. Gold/Kapton™ electrodes appear
to have always been used in an unmodified fashion. Therefore,
the quality of the gold surface after fabrication was not as critical
as when the surface is to be modified. As yet, we find that the highest quality modified electrode has been created by cleaning the
gold surface using oxygen plasma.
Cyt c was chosen for the immobilized protein after numerous
attempts to immobilize and retain the working function of SOD.
Using cited procedures [36–38], SOD can be immobilized onto a
gold surface electrostatically (3-mercaptopropionic acid, MPA) or
via DTSP. However, we found that the metal ions in active sites,
which are complexed by nearby amino acid ligands to maintain
protein shape, easily dissociated from the protein during amperometry, rendering the resultant apoenzyme useless. Since certain
thiols can yield signal in the presence of OÅÀ
2 without protein present, inactivation is typically not easily detected unless cyclic voltammetry is performed to characterize the electrodes [32].
Therefore, cyt c, while not the most selective protein for interacting
with OÅÀ
2 , is more stable than SOD when immobilized. Cyclic voltammograms of the DTSP/cyt c surface at varying scan rates can
be seen in Fig. 1. The peak potentials of the sensor near 0.05 V
and +0.1 V vs. Ag/AgCl remain stable with peak ratio of 2 indicating
a quasi reversible reaction. The xanthine/xanthine oxidase system
for producing OÅÀ

2 was used to calibrate the sensor.
The response of the sensor (3 mm in length) to multiple additions of xanthine oxidase can be seen in Fig. 2 with results summarized in Table 1, with standard deviation being that within the
single run Each addition of enzyme generates sufficient OÅÀ
2 that,
at steady state, [OÅÀ
2 ] increases by 4.5 lM. Data was processed by
boxcar averaging the signal using 3 data points (IgorPro, Wavemetrics) to reduce noise. Current was then averaged when a stable
baseline was reached and again after each addition of xanthine oxidase. Shifts in current were calculated by difference using the
background current as reference. To ensure that the signal was indeed from OÅÀ
2 , SOD was added after the four aliquots of xanthine
oxidase, dropping the current 67% from maximum towards baseline. Though, these sensors are not intended for repeated use,
SOD proved not to alter the sensing surface. Large noise fluctuations at 100, 200, 300, 400, and 500 s are due to induced current
from magnet mechanical stirring, indicating why mechanical

The overall design of the sensor kept in mind the incorporation
of a reference electrode and multiple working electrodes on the
same substrate with minimal dimensions. The overall length of
the electrode is 2.5 cm, with a tip width of 200 lm and working
length of up to 3 mm. The dimensions were governed by the opening of the round window in a Mongolian gerbil, the initial in vivo
target of this sensor, to monitor OÅÀ
2 generation in the cochlea.
Using Kapton™, the substrate is robust and the gold surface malleable enough to produce a flexible sensor. However, there are concerns with delamination of gold from the adhesion layer. Moisture
seems to be the main cause of delamination and can be minimized
by placing unmodified electrodes in a desiccator until ready for
use. After modification, gold surfaces were stable for up to 10 days
before delamination rendered the electrodes useless. Reliable resolution of the delamination issue is critical if Kapton™ is to be routinely used as a substrate. The flexing of the electrode did not
appear to cause of any visual delamination.
The heart of a perfectly modified electrode is a perfectly clean
surface. Bulk gold electrodes are typically cleaned using a variety
of processes such as mechanical polishing using alumina powder,

Fig. 1. Cyclic voltammograms of cyt c/DTSP/Au Kapton™ working electrode varying
scan rates from innermost to outermost scan: 0.1, 0.2, 0.3 and 0.4 V sÀ1 in 25 mM
phosphate buffer (pH 7.4).

and the mechanical stirring was turned on for 20 s and then turned
off. A final 50 lL injection of the SOD solution was then added to
the reaction cell to demonstrate the extent to which removal of
OÅÀ
2 returns signal to the background level. Again, mechanical stirring was employed for a brief period and then stopped while current data continued to be collected until the end of the analysis.
2.4. Interferent studies
The DTSP/cyt c electrode was rinsed with phosphate buffer and
placed in 3 mL xanthine solution (2 mM in artificial perilymph).
The sensor was then held at 200 mV vs. the reference until a stable
background was obtained. 50 lL xanthine oxidase solution was
added to the solution and the solution was mechanically stirred
for a brief time. 50 lL of interferent (final concentration of
3.75 mM) was then pipetted into the solution, mechanically stirring briefly. Finally, 50 lL SOD solution was added and mixed for
a brief time to void the solution of OÅÀ
2 .
3. Theory
The evolution of OÅÀ
2 from the reaction between xanthine and
xanthine oxidase (XO), in the presence of O2, can be seen in Eq.
(1) with the dismutation reaction of OÅÀ
2 found in Eq. (2).
½XOŠ

þ
Xanthine þ O2 þ H2 O ! Urate þ OÅÀ

2 þ 2H

ð1Þ

þ
2OÅÀ
2 þ 2H ! H2 O þ H2 O2

ð2Þ
OÅÀ
2

Assuming no interference, the concentration of
at steady
state is proportional to [XO]1/2. This presumes that dismutation,
rather than consumption of OÅÀ
2 by the electrode, is the main sink
for the radical. Presuming [O2] and [xanthine] are high enough that
xanthine oxidase concentration limits the rate of OÅÀ
2 formation,
and making the steady-state approximation,

d½OÅÀ
2
2 Š
¼ keff ½XOŠ À k2 ½OÅÀ
2 Š ¼ 0
dt

ð3Þ


1=2
keff ½XOŠ
k2

ð4Þ

½OÅÀ
2 ŠSS ¼

with keff the effective rate constant for reaction (1) and k2 the pHdependent rate constant for reaction (2).


R.C.K. Wilson et al. / Journal of Electroanalytical Chemistry 662 (2011) 100–104

Fig. 2. Chronoamperometry response (at 200 mV) of cyt c to 4 additions of 25 lL of
xanthine oxidase solution in 2 mM xanthine solution, followed by addition of
50 lLSOD (2 mg/mL) to bring signal back to background current. Data (obtained at
3 Hz) has been box averaged for every three data points.

Table 1
Calibrated response of DTSP/cyt c electrode.

lM OÅÀ
2

Average Di from baseline (pA)

Standard deviation (pA)

4.72
6.4
7.8
8.9

14
20
22
24

8
4
2
4

stirring cannot be carried out throughout the analysis. A calibration plot for the sensor can be found in Fig. 3. The linear regression
does not intersect at origin, which is not very surprising. OÅÀ
2 is dismutating, the electrode itself is consuming OÅÀ
2 , and interferents are
most likely present. The inability of the signal to return to baseline
upon the addition of SOD also hints at interference from H2O2. Recently, Kelly et al. studied the xanthine/xanthine oxidase reaction
and found H2O2 to be the major product even at short reaction
times, prior to when the long-known suicide reaction occurs
[39]. H2O2 is a known interferent when working with cyt c and
could possibly be the cause for the shift in baseline and is the basis
for studying its effects on the sensors ability to work.
It is important to note that the xanthine/xanthine oxidase system is far from ideal. The basis for using this system relies on

Fig. 3. Calibration of data collected depicted in Fig. 2. Change in current (pA) is

plotted against OÅÀ
2 concentration (lM).

103

achieving a steady state concentration of OÅÀ
2 . The xanthine/xanthine oxidase reaction deteriorates over time, causing the concentration of OÅÀ
2 to gradually drop to zero after 3 h, by which time XO
generates only H2O2 [12]. Also, xanthine oxidase activity decreases
gradually even when stored as a solid suspension. Therefore it cannot be assumed that the same supply of xanthine oxidase will create the same concentration of OÅÀ
2 from day to day. Because of this,
the xanthine oxidase activity must be calibrated spectrophotometrically using cyt c absorbance at 550.5 nm each time it is used [11].
It is not meaningful to compute standard deviations for each data
point, as it is impossible to precisely replicate concentration increments of OÅÀ
2 . Therefore, as it stands, each electrode can only be calibrated approximately by running a spectrometric analysis of the
xanthine/xanthine oxidase solution and quickly moving onto the
calibration of the cyt c electrode. Though not the intended goal,
these sensors have proven to give signal for up to five days of use.
Interferent studies focused on species anticipated to be present
during upcoming noise-induced hearing loss studies, and on species present during calibration. Uric acid and H2O2 are products
generated in the xanthine/xanthine oxidase reaction and/or a
product of OÅÀ
2 dismutation, Eq. (1) While a significant concentration of uric acid is not typical biologically, it will be present during
the calibration process until a better method for calibration (E.
Chainani, A. Keith, personal communication) of OÅÀ
2 can be perfected. NADH, citric acid and glucose are also biologically important in metabolism and are most certainly present if a cell were
to undergo apoptosis. Ascorbic acid is a common radical scavenger
and is essential for such scavenging in biological settings.
The particular interferents listed above were analyzed. A typical
interferent study can be seen in Fig. 4. Here, once the background

current became stable, 5.2 lM OÅÀ
2 (final concentration) was added
to the system. Intense noise fluctuations occur during the brief stirring followed by an increase of 100 pA. At about 300 s 3.75 mM
glucose (final concentration) was added to the system, again with
brief noise fluctuations due to mechanical stirring, resulting in a
20% drop in current. Addition of SOD allowed signal to drop 77%
back to baseline, similar to studies without interferents. The same
sensor was then used repeatedly with no further complications,
proving no biofouling from glucose. This procedure was repeated
for each interferent and a summary of results can be found in
Table 2. NADH, citric acid, and uric acid showed no significant
interference. Expected ascorbic acid scavenging abilities were
demonstrated, Table 2. Upon addition, signal dropped 60%
followed by a relatively insignificant change upon addition of
SOD. While H2O2 is known to reduce cyt c, the effects in the

Fig. 4. Chronoamperometry (at 200 mV) of DTSP/cyt c electrode (2 mM xanthine
solution) in the presence of 50 lL xanthine oxidase solution (5.2 lM OÅÀ
2 ), followed
by 50 lL of glucose (3.75 mM), and 50 lL of SOD (2 mg/mL).


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R.C.K. Wilson et al. / Journal of Electroanalytical Chemistry 662 (2011) 100–104

Table 2
Interference of common biological molecules with electrode response.
Interference

NADH
Citric acid
Uric acid
Ascorbic acid
H2O2
Glucose

Change in current from baseline (pA), (% signal drop)
XO

XO + interferent

SOD added

159
90
158
130
162
110

139 (13%)
84 (7%)
157 (1.4%)
52 (60%)
150 (8.2%)
88 (20%)

22 (86%)
36 (60%)

52.2 (67%)
39 (70%)
140 (17%)
25 (77%)

presence OÅÀ
2 are nearly immeasurable. However, H2O2 hinders the
ability of SOD to return signal to baseline. This could be due to an
exchange of OÅÀ
2 response for a H2O2 response while maintaining
the same current. Adding catalase might not clarify this issue, as
a signal could easily then be generated by OÅÀ
2 oxidation. The difference in current response between data shown in Fig. 2 versus Fig. 4
can be explained in two parts. First, the calibration of OÅÀ
2 is time
dependent, therefore the concentration of OÅÀ
2 at the time of use always has a margin of error. Second, the working area of the electrode varies as the isolated regions are hand painted with
polyimide. Future efforts are being put forth to create reproducible
working area by utilizing lithography of polyimide.
5. Conclusion
Thin film gold on Kapton™ was used to create the first flexible
OÅÀ
2 sensor for in vivo applications. The calibration showed a linear
response within the constraints established by using xanthine/xanthine oxidase as a OÅÀ
2 source. Interferent studies demonstrated
that H2O2 might prove to be a concern while other interferents
showed expected results. Further work involves incorporation a
reference electrode onto the sensor along with a sensor specific
for H2O2.
Acknowledgements

This work was supported by The National Organization for
Hearing Research Foundation, the Deafness Research Foundation,
the US Army Corps of Engineers (cooperative agreements
W911NF-07-1-0005 and W9132T-08-2-0009), the US Army Research Office (Grant W911NF-07-1-0075), and Research Corporation (Grant RA0333). We further thank Donna S. Whitlon for
drawing our attention to the correlation between NIHL and ROS.

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