An integrated imaging approach to the study of oxidative stress generation by mitochondrial dysfunction in living cells.

902 volume 118 | number 7 | July 2010 • Environmental Health Perspectives
Research
Oxidative stress is a common feature of the
mechanism of injury induced by a broad
range of environmental agents (Finkelstein
and Johnston 2004). Such environmental oxi-
dative stress can result directly from the effects
of oxidizers, electrophiles, or free radical–
generating compounds such as ozone (Steinberg
et al. 1990), quinones (Santa-Maria et al.
2005), and redox-active transition metal ions
(Valko et al. 2006). Environmental oxidative
stress can also involve the depletion of cellular
antioxidant defense mechanisms or dysregula-
tion of oxidative metabolism processes in the
cell (Ercal et al. 2001).
Analytical methods used to study oxidative
stress often focus on the detection of oxidized
biomolecules such as oxidized lipids (Kinter
1995), proteins (Kelly and Mudway 2003),
and DNA (Aust and Eveleigh 1999). Direct
detection of reactive oxygen species (ROS)
involved in cellular oxidative stress in living
cells has relied heavily on the use of the fluores-
cent indicator 2,7-dichlorodihydrofluorescein
diacetate (H2DCF-DA) (Crow 1997).
Unfortunately, the interpretation of data
obtained with this indicator has been limited
by its lack of specificity and by experimental
artifacts that include photoinstability, autoxi-
dation, and photoconversion (Marchesi et al.
1999; Rota et al. 1999).
Mitochondria are a known source of par-
tially reduced oxygen species generated as a
by-product of oxidative metabolism in the
cell (Andreyev et al. 2005). Dysregulation
of mitochondrial function with metabolic
inhibi tors has been shown to induce the
release of ROS and associated oxidative stress
(Lam et al. 2001). Toxicologically, a wide
variety of environmental contaminants rang-
ing from aromatic hydrocarbons (Senft et al.
2002) to heavy metal ions (Valko et al. 2005)
have been shown to impair mitochondrial
respiration, with ensuing production of ROS.
A number of assays measure indices of mito-
chondrial function, such as ATP concentra-
tion, citrate synthase activity, and membrane
potential (Ouhabi et al. 1998; Smiley et al.
1991), but the methodologies used to measure
mitochondrial redox potential are limited.
In the present study, we used an integrated
imaging approach to the investigation of
environmental oxidative stress resulting from
mitochondrial dysfunction. By applying estab-
lished and recently introduced indicators, this
integrated approach allows real-time measure-
ment of mitochondrial membrane potential,
hydrogen peroxide (H2O2) levels, and redox
status in living cells (Cannon and Remington
2008; Rhee 2007). A431 skin carcinoma cells
were used as representative of rapidly growing
cells with high metabolic rate and energy use
that could be inferred to have a correspond-
ingly high mitochondrial activity. Zinc sulfate
(ZnSO4) was used as a soluble form of the
zinc ion (Zn2+) that is also environmentally
relevant, because sulfate salts are known to pre-
dominate among soluble metal salts released
by combustion processes. In addition, we used
zinc pyrithione, which is used as a mildewcide
in outdoor paints, making skin cells a relevant
cell type to study with this agent.
We report that exposure to Zn2+, a
ubiquitous ambient contaminant that has
been shown to induce inflammatory (Kim
et al. 2006; Tal et al. 2006) and cytotoxic
responses (Tang et al. 2001), results in an
intracellular accumulation of H2O2 that is
associated with a decrease of mitochondrial
reducing redox potential and depolarization
Address correspondence to J.M. Samet, 104 Mason
Farm Rd., EPA Human Studies Facility, Chapel Hill,
NC 27599-7315 USA. Telephone: (919) 966-0665.
Fax: (919) 962-6271. E-mail: Samet.James@EPA.gov
*These authors contributed equally to this work.
This study was supported by a grant to C.J.C. from
the National Institutes of Health (GM 079645).
The research described in this article has been
reviewed by the National Health and Environmental
Effects Research Laboratory, U.S. Environmental
Protection Agency, and approved for publication. The
contents of this article should not be construed to rep-
resent agency policy, nor does mention of trade names
or commercial products constitute endorsement or
recommendation for use.
The authors declare they have no actual or potential
competing financial interests.
Received 11 December 2009; accepted 22 April
2010.
An Integrated Imaging Approach to the Study of Oxidative Stress Generation
by Mitochondrial Dysfunction in Living Cells
Wan-Yun Cheng,1 Haiyan Tong,2 Evan W. Miller,3 Christopher J. Chang,3 James Remington,4 Robert M. Zucker,5
Philip A. Bromberg,6 James M. Samet,2* and Thomas P.J. Hofer7*
1Department of Environmental Sciences and Engineering, University of North Carolina–Chapel Hill, Chapel Hill, North Carolina,
USA; 2Environmental Public Health Division, National Health and Environmental Effects Research Laboratory, U.S. Environmental
Protection Agency, Chapel Hill, North Carolina, USA; 3Department of Chemistry and the Howard Hughes Medical Institute, University
of California–Berkeley, Berkeley, California, USA; 4Department of Physics, Institute of Molecular Biology, University of Oregon, Eugene,
Oregon, USA; 5Toxicology Assessment Division, National Health and Environmental Effects Research Laboratory, U.S. Environmental
Protection Agency, Research Triangle Park, North Carolina, USA; 6Center for Environmental Medicine and Lung Biology, University of
North Carolina–Chapel Hill, Chapel Hill, North Carolina, USA; 7Helmholtz Zentrum München, German Research Center for Environmental
Health, Clinical Cooperation Group Inflammatory Lung Diseases, Gauting, Germany
Background: The mechanisms of action of many environmental agents commonly involve
oxidative stress resulting from mitochondrial dysfunction. Zinc is a common environmental metal-
lic contaminant that has been implicated in a variety of oxidant-dependent toxicological responses.
Unlike ions of other transition metals such as iron, copper, and vanadium, Zn2+ does not generate
reactive oxygen species (ROS) through redox cycling.
oBjective: To characterize the role of oxidative stress in zinc-induced toxicity.
Methods: We used an integrated imaging approach that employs the hydrogen peroxide
(H2O2)-specific fluorophore Peroxy Green 1 (PG1), the mitochondrial potential sensor 5,5´,6,6´-
tetrachloro-1,1´,3,3´-tetraethylbenzimidazolylcarbocyanine iodide (JC-1), and the mitochondria-
targeted form of the redox-sensitive genetically encoded fluorophore MTroGFP1 in living cells.
results: Zinc treatment in the presence of the Zn2+ ionophore pyrithione of A431 skin carcinoma
cells preloaded with the H2O2-specific indicator PG1 resulted in a significant increase in H2O2
production that could be significantly inhibited with the mitochondrial inhibitor carbonyl cyanide
3-chlorophenylhydrazone. Mitochondria were further implicated as the source of zinc-induced
H2O2 formation by the observation that exposure to zinc caused a loss of mitochondrial membrane
potential. Using MTroGFP1, we showed that zinc exposure of A431 cells induces a rapid loss of
reducing redox potential in mitochondria. We also demonstrated that zinc exposure results in rapid
swelling of mitochondria isolated from mouse hearts.
conclusion: Taken together, these findings show a disruption of mitochondrial integrity, H2O2
formation, and a shift toward positive redox potential in cells exposed to zinc. These data demon-
strate the utility of real-time, live-cell imaging to study the role of oxidative stress in toxicological
responses.
key words: biosensors, confocal microscopy, hydrogen peroxide, mitochondrial dysfunction, oxi-
dative stress, real-time imaging, ROS. Environ Health Perspect 118:902–908 (2010). doi:10.1289/
ehp.0901811 [Online 22 April 2010]

Imaging of mitochondrial oxidative stress
Environmental Health Perspectives • volume 118 | number 7 | July 2010 903
of the mitochondrial membrane. These find-
ings demonstrate the utility of an integrated
application of imaging techniques for the
study of mechanisms of environmental oxida-
tive stress in living cells with improved spatial
and temporal resolution as well as specificity.
Materials and Methods
Cell culture and experimental settings. A431
human skin carcinoma cells (no. CRL-1555;
American Type Culture Collection, Manassas,
VA, USA) were used for live cell imag-
ing experiments. The cells were cultured
in Dulbecco’s modified Eagle’s medium
(DMEM; catalog no. 11995, GIBCO,
Grand Island, NY, USA) and supplemented
with 10% fetal bovine serum (FBS) and
5 µg/mL gentamicin at 37°C, 5% CO2.
Cells were seeded on round glass cover slips
(22 mm in diameter, thickness #1) in six-well
culture plates at 150,000–250,000 cells per
well. Cultures were deprived of growth fac-
tors overnight prior to study. The A431 cells
were preloaded with Peroxy Green 1 (PG1)
or 5,5´,6,6´-tetrachloro-1,1´,3,3´-tetraethy-
lbenzimidazolylcarbocyanine iodide [JC-1
(T3168; Invitrogen, Carlsbad, CA, USA)],
and the cover slip cultures were fitted into
a custom-made stainless-steel chamber filled
with 500 µL phosphate-buffered saline (PBS),
which was supplemented with 1 g/L glu-
cose and kept at 37°C with a stage heater.
Conventional and spectral confocal micros-
copy analyses were conducted using a Nikon
Eclipse C1Si confocal microscope (Nikon
Instruments Inc., Melville, NY, USA) that was
equipped with TE 2000 microscope. Light
was delivered to the sample with a 60 × Plan
Apo 1.4 numerical aperture (NA) objective;
the system also uses diode lasers of 404 nm,
488 nm, 561 nm, and 633 nm. Prior to each
experiment, the confocal microscope was
tested for field illumination alignment, opti-
cal efficiency, colocalization, and axial resolu-
tion (Lerner and Zucker 2004; Zucker 2006a,
2006b; Zucker and Lerner 2005; Zucker et al.
2007); and the lens was inspected and cleaned
before use. Cells were exposed sequentially
to ZnSO4 (catalog no. Z-0251; Sigma, St.
Louis, MO, USA) at concentrations between
10 µM and 100 µM with or without 4 µM of
the Zn2+-specific ionophore pyrithione, given
at 5 min. The inhibitors apocynin (100 µM)
(Miller et al. 2007), wortmannin (10 µM),
diphenyleneiodonium (DPI; 25 µM) (Riganti
et al. 2004), carbonyl cyanide 3-chloro-
phenyl hydrazone (CCCP; 10 µM) (Bogeski
et al. 2006), and Ly294002 (10 µM; all
inhibitors were obtained from Sigma), plus
compound 56 (C56, 10 µM; Calbiochem,
San Diego, CA, USA) (Tal et al. 2006) and
EHT 1864 (5 µM; provided by C.J. Der,
University of North Carolina–Chapel Hill)
(Shutes et al. 2007) were applied 30 min
prior to adding Zn2+. H2O2 (1 mM), CCCP
(10 µM), or dithiothreitol (DTT; Sigma;
10 mM) was added at the end of experiments
as positive controls.
Measurement of H2O2. H2O2 produc-
tion was monitored using the fluorescein-
like Peroxy Green 1 (PG1) dye (Miller et al.
2007). PG1 is a boronate probe with high
specificity for H2O2. A431 cells grown on
glass cover slips were labeled in 5 µM PG1
for 15 min at 37°C in PBS glucose solution
prior to measurement. PG1 fluorescence was
excited using an argon laser (at λ = 488 nm),
and the emission spectrum was monitored in a
range of λ = 490 nm to 570 nm over 32 chan-
nels with 2.5 nm band pass. Signal intensity
was quantified at the PG1 emission peak at
523 nm.
Measurement of mitochondrial mem-
brane potential. The mitochondrial mem-
brane potential was monitored using the
fluorescent indicator JC-1. In the presence
of physiological mitochondrial membrane
potentials, JC-1 forms aggregates that fluo-
resce with an emission peak at 588 nm. Loss
of membrane potential favors the monomeric
form of JC-1, which has an emission peak at
530 nm. Cells were labeled with 5 µM JC-1
in DMEM supplemented with 10% FBS and
1 µg/mL gentamicin at 37°C. After 15 min
incubation, cells were washed with PBS twice
and placed in the chamber with 500 µL PBS
glucose. JC-1 fluorescence intensity was
monitored with dual excitation at 488 nm
and 561 nm and an emission scan range of
490–650 nm (32 channels, 5 nm per chan-
nel). Mitochondrial membrane potential was
inferred from the ratio of fluorescence inten-
sity of emission maximum at 593 nm and
538 nm, which represented the J-aggregate
and monomeric forms, respectively.
Cardiac mitochondrial swelling assay.
Adult pathogen-free female CD-1 mice,
which were purchased from Charles River
(Raleigh, NC, USA), were used as the source
of the cardiac mitochondria. Animals were
housed at the U.S. Environmental Protection
Agency (EPA) animal care facility (accred-
ited by the Association for Assessment and
Accreditation of Laboratory Animal Care)
and given ad libitum access to both water and
food. Animal care was given in accordance
with institutional guidelines, and animals
were treated humanely, with regard to allevi-
ating suffering. The studies were conducted
with approval by the EPA Institutional
Animal Care and Use Committee. Mice were
euthanized with an intraperitoneal injection
of sodium pentobarbital (80 mg/kg body
weight), and hearts were excised and weighed.
Freshly isolated mitochondria were prepared
from the ventricles by differential centrifuga-
tion. Briefly, heart tissues were homogenized
with three strokes of a polytron homogenizer
in ice-cold homogenization buffer contain-
ing 225 mM mannitol, 75 mM sucrose,
5 mM morpholinepropanesulfonic acid,
and 2 mM taurine, with 0.2% bovine serum
albumin (BSA; pH 7.4). The homogenate
was transferred to a glass homogenizer and
homogenized for five strokes on ice. After
centrifugation at 2,500 rpm for 5 min at
40°C, the supernatant was removed and cen-
trifuged at 8,000 rpm for 5 min. The pellet
was sequentially washed with homogenization
buffer three times and resuspended in homo-
geni zation buffer plus 5 mM KH2PO4. The
protein concentration was determined with
BSA as a standard by a Bradford assay.
The mitochondria (50 µg) were incu-
bated in buffer containing 120 mM KCl,
10 mM Tris HCl, and 5 mM KH2PO4 at
room temperature. After adding 10 mM glu-
tamate and 2 mM malate, the light scattering
of mitochondria was measured at 540 nm for
40 min with a 96-well plate spectrophotom-
eter (POLARstar Optima; BMG, Alexandria,
VA, USA). We initiated calcium- or zinc-
induced mitochondrial swelling by adding
250 µM calcium or 100 µM zinc and meas-
ured for another 20 min. The absorbance was
normalized to the initial absorbance.
Measurement of redox potential in mitochon-
dria. The genetically encoded mitochondria-
targeted form of the genetically encoded fluo-
rescent reporter redox-sensitive green fluores-
cent protein (MTroGFP1) was used for the
meas urement of redox potential in mitochon-
dria (Hanson et al. 2004). Fugene 6 (cata-
log no. 11815091001; Roche, Mannheim,
Germany) was used for transfection accord-
ing to the manufacturer’s protocol. The
MTroGFP1 plasmid was mixed with Fugene 6
for 30 min at room temperature and applied
to the A431 cells for 48 hr. Tetramethyl rho-
damine methyl ester (TMRM; catalog no.
T-668, Invitrogen), a mitochondria-specific
dye, was used to validate the transfection by
incubating 500 nM TMRM with transfected
cells for 15 min and by visualizing with exci-
tation at 561 nm and with emission filter
of 605/75 nm (Chroma Technology Corp,
Rockingham, VT, USA). Green fluorescence
was derived from excitation at both 404 nm
and 488 nm with an emission detected using
a band-pass filter of 525/50 nm (Chroma).
The results were calculated by rationing the
emissions excited by 488 nm and 404 nm
laser sequentially.
Statistical analysis. Imaging data were
collected with Nikon EZ-C1 software and
quantified by EZ-C1 and Nikon Elements.
Figures were plotted with mean ± SE, with
three repeat experiments. An average of 5–10
cells with different fluorescence intensi-
ties were collected as regions of interests in
each experiment and quantified with Nikon
EZ-C1 and Nikon Elements software (Nikon

Cheng et al.
904 volume 118 | number 7 | July 2010 • Environmental Health Perspectives
Instruments). Pairwise comparisons were car-
ried out using Student’s t-test; a p-value of
< 0.05 was considered statistically significant.
Results
Zinc-induced H2O2 production visualized by
PG1 in living cells. As a model toxicant for
these studies, we used Zn2+, a non-redox-active
metal that is ubiquitously found associated
with particulate matter in ambient air and
occupational settings. Addition of noncytotoxic
concentrations (Tal et al. 2006) of Zn2+ and
pyrithione (10–100 µM ZnSO4 plus 4 µM
pyrithione) to A431 cells resulted in a time-
dependent elevation in intracellular concen-
trations of H2O2 as detected by an increase
in PG1 fluorescence intensity (Figure 1A,B).
H2O2 added as positive control resulted in a
marked increase (550%) in PG1 fluorescence
(Figure 1C). Spectral analysis of PG1 fluores-
cence excited with 488 nm revealed an emis-
sion peak at 523 nm (Figure 1D), consistent
with published reports (Miller et al. 2007).
Sequential images were captured at 30-sec
intervals and plotted as the relative fluores-
cence intensity normalized to initial intensity.
Cells exposed to 100 µM Zn2+ for 10 min
showed a 64% increase in PG1 fluorescence
intensity relative to starting levels. PG1-loaded
resting cells not exposed to Zn2+ observed
during the same testing period showed < 4%
increase in fluorescence intensity (Figure 1E).
Similar to resting cells, A431 cells incubated
in 4 µM pyrithione alone did not show an
increase in PG1 fluorescence intensity (data
not shown).
Identification of the source of Zn2+-
induced H2O2. As shown in Figure 1, Zn2+-
stimulated H2O2 production was visualized
as an increase in fluorescence disseminated
throughout the cytosol. To identify the intra-
cellular source of H2O2, cells were pretreated
with the nico tinamide adenine dinucleotide
phosphate oxidase inhibitors apocynin or DPI,
the epidermal growth factor receptor kinase
activity inhibitor C56, the phosphoinositide
3-kinase activity inhibitors wortmannin or
Ly294002, the Rac GTPase kinase inhibitor
EHT 1864, or the mitochondrial uncoupler
CCCP, prior to exposure to Zn2+ (Bogeski
et al. 2006; Riganti et al. 2004; Shutes et al.
2007; Vanhaesebroeck et al. 2001). With the
exception of CCCP, the application of these
inhibitors did not have statistically signifi-
cant effects on Zn2+-induced H2O2 produc-
tion in A431 cells (Table 1). Pretreatment
with CCCP induced a statistically significant
32% inhibition in PG1 fluorescence intensity
relative to Zn2+ alone. Treatment with other
reagents resulted in less than 10–20% inhibi-
tion of the H2O2-dependent PG1 fluorescence
production induced by Zn2+ (Table 1). These
findings implicate mitochondria as the source
of Zn2+-induced H2O2 production.
Zinc-induced mitochondrial dysfunction.
The maintenance of the electron transport
chain proton gradient by functional mito-
chondria establishes a transmembrane electri-
cal potential that can be monitored using the
fluorescence indicator JC-1. Intrinsically a
Figure 1. Visualization of zinc-induced H2O2 production by PG1 fluorescence in A431 cells. A431 cells
were incubated with vehicle alone for 5 min (A), 100 µM zinc sulfate for 30 min (B), or 1 mM H2O2 given at
45 min (C). (D) Emission spectra confirmation of PG1 fluorescence with peak at 523 nm; intensity is shown in
arbitrary units (AU). (E) Time course of H2O2 production detected by PG1 fluorescence in resting cells and in
cells stimulated with 10 µM Zn plus 4 µM pyrithione at 5 min, 20 µM Zn at 15 min, or 70 µM Zn at 25 min; H2O2
(1 mM) was added at 35 min as a positive control for both experimental conditions. Triplicate observations
were made for control and stimulated cells with an average of 10 cells in each run. Data are mean ± SE.
600
400
200
0
Pe
rc
en
t o
f i
ni
tia
l i
nt
en
si
ty
0 10 20 30 40
Time (min)
50
20 µM Zn
70 µM Zn
1 mM H2O2
1,250
1,000
750
500
250
0
Fl
uo
re
sc
en
se
in
te
ns
ity
(A
U
)
Wavelength (nm)
480 505 530 555 580
50 µm
50 µm
50 µm
10 µM Zn +
4 µM pyrithione
Resting cells
Zn2+-stimulated cells
Table 1. The effect of inhibitors on Zn2+-induced
H2O2 production.
Inhibitora Concentration (µM) Percent inhibition
Apocynin 100 21
DPI 25 7
C56 10 3
Wortmannin 10 10
Ly294002 10 0
EHT 1864 5 5
CCCP 10 32*
aCells incubated with inhibitors in various concentrations
30 min prior to 100 µM Zn2+ exposure. Inhibition effects
were calculated by comparison of Zn2+-induced PG1
fluorescence after 30 min exposure with or without prior
inhibitor treatment.
*Denotes statistically significant difference from vehicle
control, p < 0.05.

Imaging of mitochondrial oxidative stress
Environmental Health Perspectives • volume 118 | number 7 | July 2010 905
green indicator with an emission maximum
at 529 nm in monomeric form, JC-1 accu-
mulates in functional mitochondria in con-
centrations sufficient to form J-aggregates,
which leads to a shift of the emission maxi-
mum to 588 nm (Figure 2A). Zinc-induced
mitochondrial depolarization led to a change
in the equilibrium of JC-1 observed as a shift
of the JC-1 emission maximum to a shorter
wavelength (Figure 2B), corresponding to an
emission peak shift from 538 nm to 588 nm
(Figure 2D). The ratio of the fluorescence
emission at 538 and 588 nm represents
the degree of Zn2+-induced mitochondrial
depolarization (Guthrie and Welch 2008).
Loss of mitochondrial membrane potential
was observed 10 min after cells were exposed
to 100 µM Zn2+ and 4 µM pyrithione, and
continued to rise for 30 min (Figure 2E).
CCCP was added at the end of each experi-
ment as a positive control. As shown in
Figure 2E, the addition of CCCP did not
induce a further increase in fluorescent inten-
sity in cells exposed to Zn2+, suggesting a
complete depolarization of mitochondrial
potential induced by Zn2+ in A431 cells.
As an independent measurement of mito-
chondrial function, we next examined the
effect of Zn2+ exposure on the mitochondrial
membrane transition pore using the mitochon-
drial swelling assay in isolated cardiac mouse
mitochondria. This particular assay requires
a large number of isolated mitochondria that
would be impractical to obtain from cultured
cells. Therefore, mouse heart mitochondria
were used for this purpose as a model suitable
for toxicological testing. Treatment with Zn2+
and pyrithione resulted in significant swelling
of isolated mitochondria. The addition of cal-
cium (positive control) induced spontaneous
swelling, indicated by a 17% decrease in absor-
bance at 4 min, whereas the addition of zinc
induced a similar effect, with a 15% decrease
in absorbance (Figure 3). These results inde-
pendently confirmed that Zn2+ directly affects
mitochondrial function.
Visualization of Zn2+-induced oxidative
stress in mitochondria. The data presented
above indicate that the mitochondrion
is a target of Zn2+-mediated toxicity and a
potential source of ROS and oxidative
stress. To examine the effect of Zn2+ expo-
sure on mitochondrial redox potential, we
used MTroGFP1 (Hanson et al. 2004). This
geneti cally encoded reporter responds to
changes in redox status with changes in the
relative intensity of fluores cence at 510 nm
upon excitation with its two excitation
maxima, 404 and 488 nm. Cells transfected
Figure 2. Measurement of mitochondrial membrane potential visualized by JC-1 in A431 cells treated with
zinc. A431 cells treated with vehicle alone (A) or with 100 µM zinc before (B) and after addition of 10 µM
CCCP (C) as a positive control. (D) The spectrum of JC-1 is shown under 2 different conditions; control cells
(blue line) and depolarized cells (black line). Intensity is shown in arbitrary units (AU). (E) Measurement of
JC-1 fluorescence intensity (taken as the ratio of green to red) in control and Zn2+-exposed A431 cells; 100
µM zinc plus 4 µM pyrithione were added at 5 min, and 10 µM CCCP was added to both groups at 35 min.
Images were obtained with simultaneous excitation of 488 nm and 561 nm laser and emission scan range
between 490 nm and 650 nm using a 5 nm band pass. Triplicate observations were made for control and
stimulated cells with an average of 10 cells in each run. Data are mean ± SE.
1,750
1,500
1,250
1,000
750
500
250
0
Fl
uo
re
sc
en
se
in
te
ns
ity
(A
U
)
Wavelength (nm)
500 525 550 575 625600 650
50 µm50 µm
50 µm
3.5
3.0
2.5
2.0
1.5
1.0
0.5
0
G
re
en
/r
ed
ra
tio
0 5 10 20 30 40 4515 25 35
Time (min)
100 µM Zinc +
4 µM pyrithione
10 µM CCCPZinc
Control
Figure 3. Zinc-induced mitochondrial dysfunction
measured using the swelling assay. A suspension
of isolated cardiac mitochondria was monitored
for absorbance at 550 nm after the addition of 100
µM Zn2+ or 250 µM calcium ion. Absorbance values
were normalized to the initial reading. Data repre-
sent three independent experiments.
1.1
1.0
0.9
0.8
0.7
0.6
0.5
Per
ce
nt
in
iti
al
a
bso
rba
nc
e
0 5 10
Time (min)
20 2515
100 µM Zn
250 µM Ca

Cheng et al.
906 volume 118 | number 7 | July 2010 • Environmental Health Perspectives
with MTroGFP1 displayed the expected green
fluorescence in a pattern that was exclusively
associated with mitochondria (Figure 4A).
Confirming the localization of the sensor, the
MTroGFP1 fluorescence colocalized with a
validated mitochondrial indicator, TMRM
(Figure 4B,C). Exposure of these cells to
100 µM Zn2+ and 4 µM pyrithione induced
a rapid increase in the ratio of fluorescence
at 404/488, indicating a less reduced redox
(Figure 4D). This Zn2+-induced change cor-
responded to a loss of mitochondrial reducing
potential, from a value previously reported to
be –288 mV (Hanson et al. 2004), toward a
more positive redox potential, starting within
2 min and reaching a plateau by 10 min
(Figure 4D). Subsequent addition of 10 mM
DTT as a positive control restored a negative
mitochondrial redox potential.
Discussion
Oxidative stress is increasingly recognized as an
important feature of the mechanism of toxic
action that is common to many structurally
disparate environmental contaminants (Valko
et al. 2005). However, a significant limita-
tion in the investigation of oxidative stress
in toxicology has been the availability of an
integrated methodological approach for real-
time detection of reactive oxidant species and
oxidative damage in living cells. Compared
with conventional bio chemical assays, live-cell
imaging offers superior temporal and spatial
resolution of intra cellular processes. In the
present study, we have employed an integrated
imaging approach to study oxidative stress
associated with mitochondrial dysfunction in
cells exposed to the environmental air pollu-
tant Zn2+. The detection of specific ROS is
an important feature of this approach. The
indicator H2DCF-DA and its variants have
been used widely for ROS detection by imag-
ing of living cells. However, several limitations
are associated with H2DCF-DA, including a
lack of ROS specificity and its susceptibility to
oxidation by different species such as peroxy-
nitrite, nitric oxide, superoxide, and H2O2
under various experimental conditions (Crow
1997). In addition, H2DCF-DA can also be
oxidized by heme and hemoproteins (Ohashi
et al. 2002) and is subject to photoreduction
(Marchesi et al. 1999), further reducing the
reliability of ROS detection with this indicator.
Using PG1, we visualized Zn2+-stimulated
H2O2 production in living cells. Although
PG1 is not specifically targeted to an intra-
cellular compartment, when used in combina-
tion with classical inhibitors, as we did in this
study, it can be used to infer an intracellular
source of H2O2 within the cell. PG1 con-
sists of a fluorescein-like dye conjugated to a
chemoselective boronate switch that responds
to H2O2 with high specificity (Miller et al.
2007). PG1 is the first fluorescence-based
molecular indicator for the specific detec-
tion of H2O2 with sufficient sensitivity to
detect low concentrations of peroxide such as
those produced by nonphagocytes responding
to physiological signals (Miller et al. 2007).
Thus, as an ROS sensor, PG1 represents a
significant improvement over H2DCF-DA in
that it offers superior specificity, sensitivity,
and stability (Rhee 2007). Miller et al. (2007)
showed that PG1 has high specificity for
H2O2 relative to a wide range of other oxy-
gen, nitrogen, chlorine, and organic oxidant
species. Lending credence to these findings,
in the present study we showed that Zn2+, a
transition metal incapable of producing ROS
by redox cycling, induces H2O2 production
detected by PG1 fluorescence, which we inde-
pendently show to be the result of mitochon-
drial dysfunction induced by Zn2+ exposure.
A critical element of the approach applied
in this study is monitoring redox potential
using redox-sensitive variants of green fluo-
rescent protein (roGFPs). These genetically
encoded reporters were first described by
Hanson et al. (2004) as redox potential sen-
sors with two fluorescence excitation max-
ima, thus permitting ratiometric analysis that
minimizes errors associated with variations in
indicator concentration (roGFP expression),
illumination intensity, and cell thickness. The
roGFP sensors were genetically engineered
to respond to changes in intra cellular thiol-
disulfide equilibria (Hanson et al. 2004)
and therefore provide a noninvasive method
for measuring cellular redox potential. The
roGFP sensors feature fast response rates and
selectivity for midpoint potential and for
sub cellular compartment targeting (Cannon
and Remington 2006; Dooley et al. 2004;
Lohman and Remington 2008). They have
been used for monitoring redox status in
plant cells and ischemic neuronal cells under
various conditions (Schwarzlander et al. 2009;
Vesce et al. 2005). Here, we used a mito-
chondria-targeted version, MTroGFP1, to
monitor the effect of Zn2+-induced oxidative
stress. The reporter responded as expected to
treatment with exogenous oxidants (H2O2)
Figure 4. Zinc-induced oxidative stress in mitochondria. (A) A431 cells transfected with MTroGFP1 demon-
strate green fluorescence associated with mitochondria. (B) Cells incubated with the mitochondrial indi-
cator TMRM, shown as red fluorescence. (C) Colocalization of the two images. (D) Mitochondrial redox
potential was monitored as the ratio of fluorescence intensity under 404/488 excitation normalized to the
value at 0 min; vehicle or 100 µM Zn2+ plus 4 µM pyrithione was added at 5 min. DTT (10 mM) was added
at 20 min as a positive control. Data were grouped from 20 cells studied over three separate experiments,
expressed as mean ± SE.
1.10
1.05
1.00
0.95P
er
ce
nt
o
f i
ni
tia
l r
oG
FP
1
ra
tio
0 10 155
100 µM Zn +
4 µM pyrithione
20 25 30
Time (min)
DTT 10 mM
20 µm
20 µm
20 µm
100 µM Zn2+ +
4 µM pyrithione
Vehicle